This study was supported by HKBU Seed Fund and Tier-2 Start-up Grant (RG-SGT2/18-19/SCI/009), Research Grant Council-Collaborative Research Fund (CRF-C2103-20GF) to C.H.H. Hor.

All animal-related procedures were carried out in compliance with animal handling guidelines and the protocol approved by the Department of Health, Hong Kong. Animal experiment licenses following Animal (Control of Experiments) Ordinance (Cap. 340) were obtained from the Department of Health, Hong Kong Government. The animal work was carried out in compliance with the animal safety ethics approved by HKBU Research Office and Laboratory Safety Committee.&#160;Refer to the Table of Materials for details about all materials used in this protocol.
1. Preexperiment preparation
<ol>
	<li>Preparation of culture media and buffers
	<ol>
		<li>Serum-free medium (SFM)
		<ol>
			<li>To prepare 50 mL of SFM, add 500 &#181;L of 100x L-glutamine substitute, 500 &#181;L of Penicillin-Streptomycin, 1 mM of sodium pyruvate, and 12.5 &#181;L of 1 M KCl (final 250 &#181;M) to 49 mL of Neurobasal medium.</li>
			<li>Split the SFM into 2 aliquots of 10 mL and 40 mL in 50 mL conical tubes and store them at 4 &#176;C for up to one month.</li>
		</ol>
		</li>
		<li>Digestion blocking medium: 10% FBS in SFM
		<ol>
			<li>Add 1.1 mL of heat-inactivated FBS to a 10 mL aliquot of SFM to prepare digestion blocking medium.</li>
		</ol>
		</li>
		<li>GCP culture medium: SFM with B27
		<ol>
			<li>To prepare GCP culture medium, add 800 &#181;L of serum-free B-27 supplement to a 40 mL aliquot of SFM. 
			NOTE: B-27-supplemented SFM can be stored at 4 &#176;C for up to one month. However, freshly prepared media produce optimal outcomes.</li>
		</ol>
		</li>
		<li>Dissection buffer: EBSS with glucose + HEPES
		<ol>
			<li>Add 6 g/L of glucose to calcium- and magnesium-free Earle&#39;s Balanced Salt Solution (EBSS) containing 10 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid).</li>
			<li>Sterilize the solution by passing through a 0.2 &#181;m syringe filter and store it at 4 &#176;C for long-term storage.</li>
		</ol>
		</li>
		<li>Digestion Buffer 
		NOTE: Prepare freshly before use.
		<ol>
			<li>Prepare 2 mL of digestion buffer for the digestion of 2-4 cerebellar tissues and 4 mL for 4-10 tissues. To prepare 4 mL of digestion buffer, dissolve 1.5 mg of L-cysteine (final concentration 200-400 &#181;g/mL) in 4 mL of EBSS. Invert the tube repeatedly until the powder is fully dissolved.</li>
			<li>Sterilize the solution using a 0.2 &#181;m syringe filter and a 5 mL syringe, and transfer the solution to a sterile 35 mm cell culture dish.</li>
			<li>Add 4 &#181;L of papain (1:1,000 dilution from a stock concentration of 20 units/mg) and 40 &#181;L of DNase I (1:100 dilution from a 10 mg/mL stock for a final concentration of 0.1 mg/mL).</li>
			<li>Incubate the solution at 37 &#176;C in a CO2 incubator for at least 30 min or until use. 
			&#8203;NOTE: This step is critical to activate the papain. For optimal outcomes, do not exceed 45 min at 37 &#176;C before use. Ensure the solution turns transparent and there are no white precipitates before use.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Precoating coverslips
	<ol>
		<li>To prepare the coverslips for cell attachment, incubate autoclaved 12 mm glass coverslips with 100 &#181;g/mL of poly-D-lysine (PDL, 1 mg/mL in sterile dH2O) for at least 1 h at 37 &#176;C. Keep the coverslips in the same PDL solution at 4 &#176;C until use.</li>
		<li>On the day of primary cell culture, collect the PDL in a clean conical tube and rinse the coverslips three times with sterile dH2O thoroughly to remove residual PDL. 
		NOTE: The PDL can be stored at 4 &#176;C for reuse.</li>
		<li>Transfer the PDL-coated glass coverslips onto a 24-well plate by placing one coverslip in each well.</li>
		<li>Remove excess water and add 200-300 &#181;L of Matrigel (reconstituted according to the instructions in the product datasheet in serum-free DMEM-F12) to fully cover the coverslips.</li>
		<li>Incubate the coverslips in the Matrigel for 1 h at 37 &#176;C in the CO2 incubator. 
		NOTE: Remove the Matrigel before cell seeding. This Matrigel can be collected and stored at 4 &#176;C for multiple reuses.</li>
	</ol>
	</li>
	<li>Preparation of preplating culture dish
	<ol>
		<li>Coat a 60 mm cell culture dish with 2 mL of 100 &#181;g/mL PDL (reconstitute the 1 mg/mL PDL stock solution in sterile dH2O) by incubation at 37 &#176;C for 1 h.</li>
		<li>Immediately before cell seeding, remove and collect the used PDL in a clean conical tube and store it at 4 &#176;C for multiple reuses. Rinse and wash the PDL-coated dish three times with sterile dH2O. Air-dry the culture dish before cell seeding.</li>
		<li>Use one 60 mm cell culture dish to seed cells harvested from a maximum of 2 whole-cerebellum tissues.</li>
		<li>Prepare additional culture dishes for additional cerebellums. 
		&#8203;NOTE: See steps 2.1.18 and 2.1.19 for the use of the preplating&#160;culture dish.</li>
	</ol>
	</li>
</ol>
2. Experimental day 0
<ol>
	<li>Isolation and culturing of mouse primary GCPs 
	NOTE: The GCP culture method was modified from a standard protocol, and the brief protocol was&#160;briefly described in our previous work6,11,12. For optimal GCP yield, use postnatal (P) day 6 or P7 pups for the isolation of GCP. Complete the following dissection steps 2.1.6-2.1.10 as quickly as possible for optimal cell viability.
	<ol>
		<li>Prewarm SFM, digestion blocking medium, culture medium, and Opti-MEM at 37 &#176;C during dissection.</li>
		<li>On a clean bench, presoak all the dissection apparatus in 70% ethanol for disinfection.</li>
		<li>Fill a 60 mm cell culture dish with 2-3 mL of dissection buffer and chill it on ice.</li>
		<li>Prepare fresh digestion buffer as described in section 1.1.5 and keep it warm at 37 &#176;C.</li>
		<li>Use 70% ethanol to wipe the head of the pup for disinfection.</li>
		<li>Decapitate the pup without anesthesia. Use forceps to hold the head and sterilized surgical scissors to cut from the back of the skull to decapitate the pup. Carefully remove the skin and skull to uncover the brain by using forceps.</li>
		<li>Use forceps to pinch off the cerebellum and quickly soak it in the pre-chilled dissection buffer prepared in step 2.1.3.</li>
		<li>Remove the meninges with the cerebellum submerged in the dissection buffer under a dissecting microscope. Blood vessel-enriched meninges should appear pinkish under the dissecting microscope.</li>
		<li>Remove all the meninges using forceps. 
		NOTE: A cerebellum without meninges should have a whitish appearance.</li>
		<li>Remove any visible midbrain tissues and the choroid plexus from the&#160;cerebellum.</li>
		<li>Transfer the cerebellum to a 35 mm culture dish prefilled with warm digestion buffer prepared in step 2.1.4. Avoid transferring excess dissection buffer. Cut the cerebellum into fine pieces using microspring scissors as quickly as possible.</li>
		<li>Immediately incubate the minced cerebellum at 37 &#176;C for 15 min in a CO2 incubator. Extend the incubation time to 20 min if more than 4 cerebellums are to be processed. 
		NOTE: After incubation, the tissue will clump together.</li>
		<li>Immediately transfer the digested tissue to the bottom of a new sterile 15 mL centrifuge tube using a P1000 pipette tip prewet with digestion blocking medium (avoid transferring digestion buffer).</li>
		<li>Add an appropriate volume of digestion blocking medium (1 mL for 1 cerebellum) to terminate the digestion and pipette up and down gently 30 times using a P1000 micropipette to further dissociate the tissue into a single-cell suspension. Avoid air bubble formation.</li>
		<li>Gently pass the cell suspension through a 70 &#181;m cell strainer into a new sterile centrifuge tube to remove cell clumps.</li>
		<li>Pass an additional 1 mL of fresh digestion blocking medium through the cell strainer to collect residual cells from the cell strainer.</li>
		<li>Centrifuge the filtrate at 200 &#215; g for 5 min at room temperature (RT). Remove the supernatant and resuspend the pellet with 1 mL of SFM. Repeat this step twice. Avoid forming air bubbles.</li>
		<li>Resuspend the pellet in a final volume of 2 mL of GCP culture medium and transfer the cell suspension to the PDL-coated 60 mm preplating culture dish prepared in step 1.3. Incubate for 15 min at 37 &#176;C in a CO2 incubator. 
		NOTE: Do not exceed 20 min.</li>
		<li>After incubation, tap the culture dish from the side to dislodge the loosely adherent GCP cells. Collect the adherent GCP cells in culture medium by gentle pipetting using a P1000 pipette. Collect this GCP suspension into a new 15 mL centrifuge tube. Discard the 60 mm dish. 
		NOTE: Strongly adherent astroglia and fibroblast cells will remain attached on the 60 mm dish bottom and will be separated in this step. Omitting this step will compromise the purity of the GCP culture.</li>
		<li>Count the cells and proceed to electroporation immediately.</li>
	</ol>
	</li>
	<li>Day in vitro (DIV) 0: Electroporation for Hh receptor transgene overexpression: pEGFP-Smo
	<ol>
		<li>For electroporation using a 2 mm gap cuvette, prepare the following plasmid-cell electroporation mixture for each reaction of electroporation: 1.2 &#215; 106 cells and 10 &#181;g of pEGFP-mSmo (adjust the DNA stock concentration to ~2-5 &#181;g/&#181;L in Tris-EDTA buffer or sterile dH2O) in 100 &#181;L of Opti-MEM. 
		NOTE: Reduce the total cell number if the cells are insufficient (though this may reduce the electroporation efficiency). However, do not adjust the amount of plasmid nor the total volume of Opti-MEM used per cuvette. If the total cell number required for the experiment exceeds 1.5 &#215; 106&#160;cells, scale up the electroporation mixture accordingly and perform multiple electroporation reactions separately. The cuvette may be reused up to 5 times.</li>
		<li>Prepare the post-electroporation cell seeding plate by adding 0.5 mL of culture medium into each well of the 24-well culture plate containing coated coverslips (from step 1.2) and keep it warm at 37 &#176;C in a CO2 incubator.</li>
		<li>From step 2.1.20, pipette the required number of cells into a sterile 1.5 mL microcentrifuge tube and spin at 200 &#215; g&#160;for 5 min at RT. Discard the supernatant and resuspend the cell pellet in 200 &#181;L of Opti-MEM. Repeat this step twice with Opti-MEM to ensure no residual culture medium is present in the tube 
		NOTE: For every well/coverslip of a 24-well plate, electroporate and seed 1.2-1.3 &#215; 106 cells/well to obtain ~70-75% cell confluency on the next day of culture.</li>
		<li>Set the parameters of electroporation as shown in Table 1.</li>
		<li>Pipette the electroporation reaction gently to mix well and use a long P200 tip to transfer an exact volume of 100 &#181;L of the mixture into the 2 mm gap cuvette. Avoid forming bubbles.</li>
		<li>Place the cuvette in the cuvette chamber.</li>
		<li>Press the &#937; button of the electroporator (see the Table of Materials) and make a note of the impedance value, which should be ~30-35. To ensure the electric impedance value &#937; falls within the range of 30-35, adhere to a precise volume of 100 &#181;L.</li>
		<li>Press the start button to initiate the pulse.</li>
		<li>Record the values of the measured current and joules shown on the reading frame.</li>
		<li>Remove the cuvette from the cuvette chamber.</li>
		<li>Immediately add 100 &#181;L of prewarmed culture medium into the cuvette and resuspend it by gentle pipetting up and down 2-3 times. Immediately transfer the cell suspension to the 24-well plate prepared in step 2.2.2. 
		NOTE: To minimize cell death, seed the cells immediately after electroporation.</li>
		<li>Incubate the cells at 37 &#176;C in a CO2 incubator. Leave the cells undisturbed for 3 h to avoid cell detachment.</li>
		<li>At 3 h post seeding, gently aspirate and discard half of the supernatant medium to remove floating dead cells and debris and replace with the same amount of prewarmed culture medium.</li>
		<li>Incubate the cells at 37 &#176;C&#160;in the CO2 incubator and replenish half of the medium every other day.</li>
		<li>Observe the cells under the fluorescence microscope on the next day, i.e., DIV1 (Figure 1) for GFP signal to determine the efficiency of Smo overexpression.</li>
		<li>For the stimulation of the Hh signaling pathway on DIV 1 after replenishing half of the medium, add 0.2 &#181;M Smoothened agonist (SAG) to the cells while adding an equal volume of DMSO to the control. Incubate for 24 h prior to cell fixation. 
		&#8203;NOTE: Keep the volume of DMSO/SAG added as low as possible. Here, 0.5 &#181;L DMSO or 0.5 &#181;L of 0.2 mM SAG was added per 500 &#181;L of medium per well, which is a dilution ratio of 1:1,000.</li>
	</ol>
	</li>
</ol>
3. DIV 2: Visualization of primary cilia and investigation of the Hh signaling pathway
<ol>
	<li>Cell fixation
	<ol>
		<li>At 24 h post treatment, remove the culture medium and rinse the cells 2-3 times with phosphate-buffered saline (PBS) using a disposable Pasteur pipette while&#160;avoid dislodging the cells.</li>
		<li>Fix the cells by adding ~400 &#181;L of 4% paraformaldehyde (prepared in PBS) and incubate at RT for 10 min.</li>
		<li>Rinse and wash 2-3 times with PBS using a disposable Pasteur pipette.</li>
		<li>Store in PBS at 4 &#176;C for up to 2 months or proceed to immunostaining.</li>
	</ol>
	</li>
	<li>Immunocytochemical staining of GCPs and primary cilium marker
	<ol>
		<li>In a 24-well plate, washthe cells twice in PBS with gentle shaking for 5 min each time.</li>
		<li>Add 0.5 mL of 100 mM ammonium chloride to the cell and incubate at RT for 10 min to quench the fixative.</li>
		<li>Rinse once and wash 2-3 times with PBS with gentle shaking for 10 min each time.</li>
		<li>Gently pipette 30 &#181;L of blocking buffer (BB: 0.1% Triton X-100, 1% BSA, 2% heat-inactivated horse serum in PBS) onto a piece of parafilm to form a droplet without air bubbles.</li>
		<li>Gently transfer the coverslip from the well onto the BB droplet with the cell-seeded side facing downwards. Incubate the cells with BB in a humid chamber for 1 h at RT.</li>
		<li>Prepare the primary antibody mix in BB as shown in the Table of Materials. To probe the cells with primary antibody, repeat steps 3.2.4 and 3.2.5, replacing BB with the primary antibody mix. Incubate the cells with the primary antibody for 2 h at RT.</li>
		<li>Using forceps, transfer the coverslips back to the 24-well plate with the cell-seeded side facing upwards. Rinse the cells once and wash 3-4 times in PBS with gentle shaking for 10 min each time.</li>
		<li>Prepare the secondary antibody mix in BB as shown in the Table of Materials. To incubate the cells with the secondary antibody, repeat steps 3.2.4 and 3.2.5, replacing BB with the secondary antibody mix. Incubate in the dark for 1 h at RT.</li>
		<li>Repeat step 3.2.7 for post-secondary antibody incubation washing.</li>
		<li>Mount the coverslip on a clean microscopic glass slide using mounting medium.</li>
		<li>Air-dry the slide in the dark overnight and proceed to confocal imaging13.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Chang, C. 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=Atoh1+controls+primary+cilia+formation+to+allow+for+SHH-triggered+granule+neuron+progenitor+proliferation.">Atoh1 controls primary cilia formation to allow for SHH-triggered granule neuron progenitor proliferation.</a> Developmental Cell. 48 (2), 184-199 (2019).</li><li>Dahmane, N., Ruizi Altaba, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sonic+Hedgehog+regulates+the+growth+and+patterning+of+the+cerebellum.">Sonic Hedgehog regulates the growth and patterning of the cerebellum.</a> Development. 126 (14), 3089-3100 (1999).</li><li>Lewis, P. M., Gritti-Linde, A., Smeyne, R., Kottmann, A., Mcmahon, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sonic+Hedgehog+signaling+is+required+for+expansion+of+granule+neuron+precursors+and+patterning+of+the+mouse+cerebellum.">Sonic Hedgehog signaling is required for expansion of granule neuron precursors and patterning of the mouse cerebellum.</a> Developmental Biology. 270 (2), 393-410 (2004).</li><li>Wechsler-Reya, R. J., Scott, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+neuronal+precursor+proliferation+in+the+cerebellum+by+Sonic+Hedgehog.">Control of neuronal precursor proliferation in the cerebellum by Sonic Hedgehog.</a> Neuron. 22 (1), 103-114 (1999).</li><li>Bangs, F., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+cilia+and+mammalian+Hedgehog+signaling.">Primary cilia and mammalian Hedgehog signaling.</a> Cold Spring Harbor Perspectives in Biology. 9 (5), 028175 (2017).</li><li>Hor, C. H. H., Lo, J. C. W., Cham, A. L. S., Leung, W. Y., Goh, E. L. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multifaceted+functions+of+Rab23+on+primary+cilium-+and+Hedgehog+signaling-mediated+cerebellar+granule+cell+proliferation.">Multifaceted functions of Rab23 on primary cilium- and Hedgehog signaling-mediated cerebellar granule cell proliferation.</a> Journal of Neuroscience. 41 (32), 6850-6863 (2021).</li><li>Han, Y. G., 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=Dual+and+opposing+roles+of+primary+cilia+in+medulloblastoma+development.">Dual and opposing roles of primary cilia in medulloblastoma development.</a> Nature Medicine. 15 (9), 1062-1065 (2009).</li><li>Spassky, 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=Primary+cilia+are+required+for+cerebellar+development+and+Shh-dependent+expansion+of+progenitor+pool.">Primary cilia are required for cerebellar development and Shh-dependent expansion of progenitor pool.</a> Developmental Biology. 317 (1), 246-259 (2008).</li><li>Wang, T., Larcher, L., Ma, L., Veedu, R. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systematic+screening+of+commonly+used+commercial+transfection+reagents+towards+efficient+transfection+of+single-stranded+oligonucleotides.">Systematic screening of commonly used commercial transfection reagents towards efficient transfection of single-stranded oligonucleotides.</a> Molecules. 23 (10), 2564 (2018).</li><li>Chicaybam, 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=An+efficient+electroporation+protocol+for+the+genetic+modification+of+mammalian+cells.">An efficient electroporation protocol for the genetic modification of mammalian cells.</a> Frontiers in Bioengineering and Biotechnology. 4, 99 (2017).</li><li>Lee, H. Y., Greene, L. A., Mason, C. A., Chiara Manzini, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+culture+of+post-natal+mouse+cerebellar+granule+neuron+progenitor+cells+and+neurons.">Isolation and culture of post-natal mouse cerebellar granule neuron progenitor cells and neurons.</a> Journal of Visualized Experiments: JoVE. (23), e990 (2009).</li><li>Mizoguchi, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+cerebellar+development+in+mice+overexpressing+VGF.">Impaired cerebellar development in mice overexpressing VGF.</a> Neurochemical Research. 44 (2), 374-387 (2019).</li><li>Zhou, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Confocal+imaging+of+nerve+cells.">Confocal imaging of nerve cells.</a> Current Laboratory Methods in Neuroscience Research. , 235-247 (2014).</li><li>Corbit, K. 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=Vertebrate+Smoothened+functions+at+the+primary+cilium.">Vertebrate Smoothened functions at the primary cilium.</a> Nature. 437 (7061), 1018-1021 (2005).</li></ol>

The authors have no conflicts of interest to declare.

Transfection of transgenes in primary GCP culture by electroporation method is typically associated with low cell viability and poor transfection efficiency9,10. This paper introduces a cost-effective and reproducible electroporation protocol that has demonstrated high efficiency and viability. In addition, we also demonstrate a straightforward method of studying the primary cilium-dependent Hh signaling pathway in primary GCP cells.
O...

Cerebellar GCPs are widely used to study the machinery of the Hh signaling pathway in neuronal progenitor cell-types owing to their high abundance and high sensitivity to the Hh signaling pathway in vivo1,2,3,4. In GCPs, the primary cilium acts as a pivotal Hh signal transduction hub5 that orchestrates the proliferation of the precursor cells6,7,8. In vitro visualization of Hh signaling components on the primary cilium is often challenging due to their low endogenous basal levels. Hence, transgene modification of protein expression levels and fluorophore tagging of the gene of interest are useful approaches to study the pathway at molecular resolution. However, genetic manipulation of GCP primary cultures using liposome-based transfection approaches often result in low transfection efficiency, hindering further molecular investigations9. Electroporation increases the efficiency but commonly requires exorbitant vendor-specific and cell type-restricted electroporation reagents10.
This paper introduces a high-efficiency and cost-effective electroporation method to manipulate the Hh signaling pathway components in GCP primary cultures. Using this modified electroporation protocol, a green fluorescent protein (GFP)-tagged Smoothened transgene (pEGFP-Smo) was efficiently delivered to GCPs and achieved high cell survival and transfection rates (80-90%). Furthermore, as evidenced by the immunocytochemical staining, the transfected GCPs showed high sensitivity to Smoothened agonist-induced activation of the Hh signaling pathway by trafficking EGFP-Smo to the primary cilia. This protocol shall be directly applicable and beneficial for experiments that involve in vitro genetic modification of cell types that are difficult to transfect, such as human and rodent primary cell cultures, as well as human induced&#160;pluripotent stem cells.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>GCP Culture</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>B27 supplement</td>
			<td>Life Technologies LTD</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell strainer, 70 &#181;m</td>
			<td>Corning</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I from bovine pancreas</td>
			<td>Roche</td>
			<td>11284932001</td>
			<td></td>
		</tr>
		<tr>
			<td>Earle&#8217;s Balanced Salt Solution</td>
			<td>Gibco, Life Technologies</td>
			<td>14155063</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS, qualified</td>
			<td>Thermo Scientific</td>
			<td>SH30028.02</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutamMAXTM-I ,100x</td>
			<td>Gibco, Life Technologies</td>
			<td>35050061</td>
			<td>L-glutamine substitute</td>
		</tr>
		<tr>
			<td>L-cysteine</td>
			<td>Sigma Aldrich</td>
			<td>C7352</td>
			<td></td>
		</tr>
		<tr>
			<td>Matrigel</td>
			<td>BD Biosciences</td>
			<td>354277</td>
			<td>Basement membrane matrix</td>
		</tr>
		<tr>
			<td>Neurobasal</td>
			<td>Gibco, Life Technologies</td>
			<td>21103049</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain,suspension</td>
			<td>Worthington Biochemical Corporation</td>
			<td>LS003126</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-lysine Hydrobromide</td>
			<td>Sigma Aldrich</td>
			<td>P6407</td>
			<td></td>
		</tr>
		<tr>
			<td>SAG</td>
			<td>Cayman Chemical</td>
			<td>11914-1</td>
			<td>Smoothened agonist</td>
		</tr>
		<tr>
			<td>IF staining</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma Aldrich</td>
			<td>A7906</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma Aldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma Aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
		<tr>
			<td>Primary antibody mix</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-GFP-goat ab</td>
			<td>Rockland</td>
			<td>600-101-215</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Anti-Arl13b mouse monoclonal ab</td>
			<td>NeuroMab</td>
			<td>75-287</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Anti-Pax6 rabbit polyclonal ab</td>
			<td>Covance</td>
			<td>PRB-278P</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Secondary antibody mix</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 donkey anti-goat IgG</td>
			<td>Invitrogen</td>
			<td>A-11055</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Alexa Fluor 555 donkey anti-mouse IgG</td>
			<td>Invitrogen</td>
			<td>A-31570</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 donkey anti-rabbit IgG</td>
			<td>Invitrogen</td>
			<td>A-31573</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Thermo Scientific</td>
			<td>62247</td>
			<td>Dilution Factor: 1 : 1000</td>
		</tr>
		<tr>
			<td>Electroporation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CU 500 cuvette chamber</td>
			<td>Nepagene</td>
			<td>CU500</td>
			<td></td>
		</tr>
		<tr>
			<td>EPA Electroporation cuvette (2 mm gap)</td>
			<td>Nepagene</td>
			<td>EC-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM</td>
			<td>Life Technologies LTD</td>
			<td>31985070</td>
			<td>reduced-serum medium for transfection</td>
		</tr>
		<tr>
			<td>pEGFP-mSmo</td>
			<td>Addgene</td>
			<td>25395</td>
			<td></td>
		</tr>
		<tr>
			<td>Super Electroporator NEPA21 TYPE II In Vitro and In Vivo Electroporation</td>
			<td>Nepagene</td>
			<td>NEPA21</td>
			<td>electroporator</td>
		</tr>
	</tbody>
</table>

efficient and cost effective electroporation method to study primary cilium-dependent signaling pathways in the granule cell precursor

The primary cilium is a critical signaling organelle found on nearly every cell that transduces Hedgehog (Hh) signaling stimuli from the cell surface. In the granule cell precursor (GCP), the primary cilium acts as a pivotal signaling center that orchestrates precursor cell proliferation by modulating the Hh signaling pathway. The investigation of primary cilium-dependent Hh signaling machinery is facilitated by in vitro genetic manipulation of the pathway components to visualize their dynamic localization to the primary cilium. However, transfection of transgenes in the primary cultures of GCPs using the currently known electroporation methods is generally costly and often results in low cell viability and undesirable transfection efficiency. This paper introduces an efficient, cost-effective, and simple electroporation protocol that demonstrates a high transfection efficiency of ~80-90% and optimal cell viability. This is a simple, reproducible, and efficient genetic modification method that is applicable to the study of the primary cilium-dependent Hedgehog signaling pathway in primary GCP cultures.

Using the Opti-MEM (see the Table of Materials) as the universal reagent, this proposed electroporation methodology could achieve consistently high electroporation efficiency at ~80-90% (Figure 1). The electroporation efficiency of the Smo-EGFP vector was determined at DIV 2 post electroporation by quantification of the percentage of green fluorescence-positive cells in all paired box protein-6 (Pax6)-expressing GCP cells. The electroporation efficiency of DMSO- and SAG-trea...

Watch this Scientific Journal Video about Efficient and Cost Effective Electroporation Method to Study Primary Cilium-Dependent Signaling Pathways in the Granule Cell Precursor at JoVE.com

Efficient and Cost Effective Electroporation Method to Study Primary Cilium-Dependent Signaling Pathways in the Granule Cell Precursor

Here, we present a reproducible in vitro electroporation protocol for genetic manipulation of primary cerebellar granule cell precursors (GCPs) that is cost-effective, efficient, and viable. Moreover, this protocol also demonstrates a straightforward method for the molecular study of primary cilium-dependent Hedgehog signaling pathways in primary GCP cells.

The primary cilium is a critical signaling organelle found on nearly every cell that transduces Hedgehog (Hh) signaling stimuli from the cell surface. In ...

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2.2K Views.  Hong Kong Baptist University. Here, we present a reproducible in vitro electroporation protocol for genetic manipulation of primary cerebellar granule cell precursors (GCPs) that is cost-effective, efficient, and viable. Moreover, this protocol also demonstrates a straightforward method for the molecular study of primary cilium-dependent Hedgehog signaling pathways in primary GCP cells.

Video: Efficient and Cost Effective Electroporation Method to Study Primary Cilium-Dependent Signaling Pathways in the Granule Cell Precursor

Design and Construction of a Cost Effective Headstage for Simultaneous Neural Stimulation and Recording in the Water Maze

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial products are often expensive, not easily customized, and not submersible. Here we describe a method to design and build a customized, integrated circuit headstage for simultaneous 4-channel neural recording and 2-channel simulation in awake, behaving animals. The headstage is designed using a free, commercially available CAD-type design package, and can be modified easily to accommodate different scales (e.g. to add channels). A customized printed circuit board is built using surface mount resistors, capacitors and operational amplifiers to construct the unity gain source follower circuit. The headstage is made water-proof with a combination of epoxy, parafilm and a synthetic rubber putty. We have successfully used this device to record local field potentials and stimulate different brain regions simultaneously via independent channels in rats swimming in a water maze. The total cost is &lt; $30/unit and can be manufactured readily.

We present a low-cost method to design and construct a light headstage pre-amplifier system with simultaneous neural recording and stimulation capability. This device can be waterproofed for use in swimming animals.

Headstage preamplifiers and source followers are commonly used to study neural activity in behavioral neurophysiology experiments. Available commercial ...

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental pathways. Several powerful and well established transgenic technologies are available to manipulate gene expression levels in mouse, allowing for the generation of both loss- and gain-of-function models. However, the generation of mouse transgenics is both costly and time consuming. Alternative methods of gene manipulation have therefore been widely sought. In utero electroporation is a method of gene delivery into live mouse embryos1,2 that we have successfully adapted3,4. It is largely based on the success of in ovo electroporation technologies that are commonly used in chick5. Briefly, DNA is injected into the open ventricles of the developing brain and the application of an electrical current causes the formation of transient pores in cell membranes, allowing for the uptake of DNA into the cell. In our hands, embryos can be efficiently electroporated as early as embryonic day (E) 11.5, while the targeting of younger embryos would require an ultrasound-guided microinjection protocol, as previously described6. Conversely, E15.5 is the latest stage we can easily electroporate, due to the onset of parietal and frontal bone differentiation, which hampers microinjection into the brain. In contrast, the retina is accessible through the end of embryogenesis. Embryos can be collected at any time point throughout the embryonic or early postnatal period. Injection of a reporter construct facilitates the identification of transfected cells.
To date, in utero electroporation has been most widely used for the analysis of neocortical development1,2,3,4. More recent studies have targeted the embryonic retina7,8,9 and thalamus10,11,12. Here, we present a modified in utero electroporation protocol that can be easily adapted to target different domains of the embryonic CNS. We provide evidence that by using this technique, we can target the embryonic telencephalon, diencephalon and retina. Representative results are presented, first showing the use of this technique to introduce DNA expression constructs into the lateral ventricles, allowing us to monitor progenitor maturation, differentiation and migration in the embryonic telencephalon. We also show that this technique can be used to target DNA to the diencephalic territories surrounding the 3rd ventricle, allowing the migratory routes of differentiating neurons into diencephalic nuclei to be monitored. Finally, we show that the use of micromanipulators allows us to accurately introduce DNA constructs into small target areas, including the subretinal space, allowing us to analyse the effects of manipulating gene expression on retinal development.

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

In utero electroporation allows for rapid gene delivery in a spatially- and temporally-controlled manner in the developing central nervous system (CNS). Here we describe a highly adaptable in utero electroporation protocol that can be used to deliver expression constructs into multiple embryonic CNS domains, including the telencephalon, diencephalon and retina.

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...

Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms responsible for those differences remain unclear. Primary disassociated embryonic neuronal culture provides a simplified experimental model with which to investigate the neuronal cell signaling involved in cell death as a result of ischemia or disease; however, most neuronal cultures used in research today are mixed sex. Researchers can and do test the effects of sex steroid treatment in mixed sex neuronal cultures in models of neuronal injury and disease, but accumulating evidence suggests that the female brain responds to androgens, estrogens, and progesterone differently than the male brain. Furthermore, neonate male and female rodents respond differently to ischemic injury, with males experiencing greater injury following cerebral ischemia than females. Thus, mixed sex neuronal cultures might obscure and confound the experimental results; important information might be missed. For this reason, the Herson Lab at the University of Colorado School of Medicine routinely prepares sex-stratified primary disassociated embryonic neuronal cultures from both hippocampus and cortex. Embryos are sexed before harvesting of brain tissue and male and female tissue are disassociated separately, plated separately, and maintained separately. Using this method, the Herson Lab has demonstrated a male-specific role for the ion channel TRPM2 in ischemic cell death. In this manuscript, we share and discuss our protocol for sexing embryonic mice and preparing sex-stratified hippocampal primary disassociated neuron cultures. This method can be adapted to prepare sex-stratified cortical cultures and the method for embryo sexing can be used in conjunction with other protocols for any study in which sex is thought to be an important determinant of outcome.

Primary disassociated embryonic hippocampal neuronal cultures are useful for investigating the signaling mechanisms involved in neuron death. Sexing the embryos before the isolation and dissociation of the hippocampus allows the preparation of separate male and female cultures, which enables the researcher to identify and investigate sex-specific cell signaling.

Sex differences in neuronal susceptibility to ischemic injury and neurodegenerative disease have long been observed, but the signaling mechanisms ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

FIM Imaging and FIMtrack: Two New Tools Allowing High-throughput and Cost Effective Locomotion Analysis

The analysis of neuronal network function requires a reliable measurement of behavioral traits. Since the behavior of freely moving animals is variable to a certain degree, many animals have to be analyzed, to obtain statistically significant data. This in turn requires a computer assisted automated quantification of locomotion patterns. To obtain high contrast images of almost translucent and small moving objects, a novel imaging technique based on frustrated total internal reflection called FIM was developed. In this setup, animals are only illuminated with infrared light at the very specific position of contact with the underlying crawling surface. This methodology results in very high contrast images. Subsequently, these high contrast images are processed using established contour tracking algorithms. Based on this, we developed the FIMTrack software, which serves to extract a number of features needed to quantitatively describe a large variety of locomotion characteristics. During the development of this software package, we focused our efforts on an open source architecture allowing the easy addition of further modules. The program operates platform independent and is accompanied by an intuitive GUI guiding the user through data analysis. All locomotion parameter values are given in form of csv files allowing further data analyses. In addition, a Results Viewer integrated into the tracking software provides the opportunity to interactively review and adjust the output, as might be needed during stimulus integration. The power of FIM and FIMTrack is demonstrated by studying the locomotion of Drosophila larvae.

FIM is a novel, cost effective imaging system designed to track small moving objects such as C. elegans, planaria or Drosophila larvae. The accompanying FIMTrack program is designed to deliver fast and efficient data analysis. Together, these tools allow high-throughput analysis of behavioral traits.

The analysis of neuronal network function requires a reliable measurement of behavioral traits. Since the behavior of freely moving animals is variable to ...

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, their distinct dendritic morphologies, their specific patterns of axonal connectivity, and their selective firing responses. The molecular and cellular mechanisms responsible for these aspects of differentiation during development are still poorly understood.
Here, we describe combined protocols for labeling and characterizing the structural connectivity and excitability of cortical neurons. Modification of the in utero electroporation (IUE) protocol allows the labeling of a sparse population of neurons. This, in turn, enables the identification and tracking of the dendrites and axons of individual neurons, the precise characterization of the laminar location of axonal projections, and morphometric analysis. IUE can also be used to investigate changes in the excitability of wild-type (WT) or genetically modified neurons by combining it with whole-cell recording from acute slices of electroporated brains. These two techniques contribute to a better understanding of the coupling of structural and functional connectivity and of the molecular mechanisms controlling neuronal diversity during development. These developmental processes have important implications on axonal wiring, the functional diversity of neurons, and the biology of cognitive disorders.

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

This manuscript provides protocols that use in utero electroporation (IUE) to describe the structural connectivity of neurons at the single-cell level and the excitability of fluorescently labeled neurons. Histology is used to characterize dendritic and axonal projections. Whole-cell recording in acute slices is used to investigate excitability.

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...