Work supported in part by the Walter S. and Lucienne Driskill Foundation and NIH R01NS058792.

All procedures in this protocol were conducted according to the National Institute of Health guidelines for the ethical treatment and care of animals, and according to an approved IACUC animal use protocol.
1. Preparation of Tissue Culture Dishes, Culture Media, and Dissection Instruments to be Performed at Least One Day Before Dissection
<ol>
	<li>Prepare 1 L borate buffer: 3.1 g/L boric acid and 4.75 g/L Borax in double distilled H2O. Adjust pH to 8.4. Filter-sterilize and store at 4 &#176;C until use.</li>
	<li>Dilute poly-D-lysine to 0.1 mg/ml.
	<ol>
		<li>Under a laminar-flow tissue culture hood, place enough diluted poly-D-lysine to generously cover the bottom of each well of a multi-well, culture-treated, sterile tissue culture plate. (If desired, precleaned, sterile, glass-cover slips can be added to the bottom of each well before the addition of poly-D-lysine. Cover slips can float after the addition of poly-D-lysine, so they should be pushed down with sterile forceps if necessary.)</li>
		<li>Incubate the plates overnight in the incubator at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Prepare 1 L of dissection buffer: 350 mg/L NaHCO3, 2.38 g/L HEPES, 1.44 g/L MgSO4, and 300 mg/L BSA in 1x Hanks Balanced Salt Solution without Ca, Mg, sodium pyruvate, or phenol red. Adjust pH to 7.3. Filter-sterilize and store at 4 &#176;C until needed.</li>
	<li>Prepare 500 ml plating medium: 10% Fetal Bovine Serum, 100 units/ml each of penicillin and streptomycin, and 2.5 mM glutamine in MEM without phenol red. Filter-sterilize. Store at 4 &#176;C until needed. Discard any unused medium after 1 month.</li>
	<li>Prepare 500 ml feeding medium: 2% B-27 supplement, 100 units/ml each of penicillin and streptomycin, and 2.0 mM GlutaMAX&#160;(not glutamine) in Neurobasal without phenol red. Filter-sterilize. Store at 4 &#176;C until needed. Discard any remaining feeding medium after 1 month.</li>
	<li>Prepare 10 ml of 100 mM CaCl2, 500 mM EDTA, and 1 M&#160;NaOH as needed according to standard methods.</li>
	<li>Autoclave the surgical/dissection instruments and store under sterile conditions until needed. This procedure requires two sharp extra-fine tipped angled forceps, a fine tipped curved forceps, a serrated curved forceps, a small rat-toothed forceps, one pair of micro-surgical scissors, one pair of fine, narrow tipped surgical scissors, one pair of medium tipped scissors, and a blunt-ended dissection probe. Additional sterile instruments can be added according to personal preference.</li>
</ol>
2. Preparation of Tissue Culture Dishes, Culture Media, Dissection Instruments, and Dissection Area to be Performed on the Day of Dissection
<ol>
	<li>Wash the poly-D-lysine coated tissue culture plates. Under the laminar flow biosafety cabinet, aspirate the poly-D-lysine with a vacuum line or pipette, add water to each well, and incubate at room temperature for ~5 min. Aspirate the water and replace it with clean, sterile water. Repeat 3x.
	<ol>
		<li>Allow the plates to fully dry at room temperature.</li>
	</ol>
	</li>
	<li>Prepare the dissection area by wiping down the counter and dissection scope with 70% ethanol.
	<ol>
		<li>Saturate a large Kimwipe with 70% ethanol, and lay out the autoclaved dissection tools. Cover with another Kimwipe.</li>
		<li>Fill a large, flat dish with ice. Label and fill an appropriate number of 60 mm culture dishes with cold dissection solution. Prepare enough dishes to keep the male and female heads, brains, and hippocampi separate (if applicable).</li>
		<li>Fill two larger 100 mm culture dishes with cold dissection buffer.</li>
		<li>Set aside a separate 100 mm dish without buffer (dry).</li>
		<li>Establish an area for biohazard waste and another for animal tissue (to be disposed of appropriately after the dissection, according to institutional standards).</li>
	</ol>
	</li>
	<li>Warm 30 ml plating media in a 37 &#176;C water bath. This warm solution will be used in the cell disassociation phase of the protocol.</li>
	<li>Place 1 ml of plating media into each well of 24-well, poly-D-lysine coated, tissue culture plate. Use an appropriate smaller volume if smaller plates are used. Place the covered plate into a 37 &#176;C incubator at 5% CO2 to warm and equilibrate to the CO2 during the dissection.</li>
	<li>Prepare the papain solution.
	<ol>
		<li>Label two 15 ml sterile conical tubes, one as &#34;male&#34;, and the other as &#34;female&#34;.</li>
		<li>Add the following to each tube: 10 ml of dissection buffer, 150 &#956;l 100 mM CaCl2, 10 &#956;l 500 mM EDTA, 10&#956;l 1 M NaOH, and 150 U of papain.</li>
		<li>Gently mix by inverting the tube three times, then warm in a 37 &#176;C water bath. Note: Each vial of papain will have a different unit of activity per mg of papain, thus the volume of papain added to solution will differ from vial to vial. The papain will be cloudy initially, but the solution will clear as the papain dissolves completely.</li>
	</ol>
	</li>
</ol>
3. Harvesting, Sexing, and Dissection of E18 Embryos
<ol>
	<li>Anesthetize the pregnant female mouse with isoflurane or other appropriate anesthesia. Euthanize the mouse under deep anesthesia by decapitation or cervical dislocation.</li>
	<li>Working quickly, clean the female&#39;s abdomen with 70% ethanol. If desired, the abdomen can be shaved before being sprayed with ethanol to reduce risks of contamination. Lift the abdominal skin with the rat-toothed forceps. Make a large incision in the abdomen with the medium-tipped scissors, cutting through skin and abdominal muscles, exposing the internal organs.</li>
	<li>Carefully remove the embryos, lifting each entire uterine horn clear of the abdomen, being careful not to let it contact any external tissue if possible (to avoid contamination). Place the uterine horns containing the embryos into a dish with cold dissection saline and transfer to ice.</li>
	<li>Carefully remove each pup from the uterine horns and the embryonic sac. The pups are fragile, so this is best done by cutting the uterine tissue between each embryo to expose the embryonic sac, and then carefully slicing the sac on the surface with the edge of the small, narrow scissors. The embryonic fluid and the pup can then be gently pushed through the opening without damage to the embryo.
	<ol>
		<li>If the incision in the embryonic sac is not large enough for the fluid and pup to come out easily, then enlarge the incision carefully until the pup slides out with minimal force.</li>
		<li>Do not squeeze the sac too forcefully to force the embryo out, as this too could damage the embryos.</li>
	</ol>
	</li>
	<li>Remove the head from a single embryo. Leave the head in a dish of dissection saline on ice. Place the body of the embryo on its back on the dry 100 mM dish under the dissection scope. Using the curved medium-tipped forceps in one hand and the serrated curved forceps in the other, gently flatten the pup out, pushing the back legs down and ventral, the front legs down and dorsal.</li>
	<li>Remove the tail with scissors if necessary; the tail tends to curl up over the abdomen and can get in the way of the dissection.</li>
	<li>Release the back legs and pinch the abdomen at the root of the umbilical cord, called the umbilicus. Lift up gently. Release the front legs and with the other hand and the micro-scissors, make a small inverted &#34;v&#34; in the abdomen just above the umbilicus with the point of the &#34;v&#34; pointing cranial.</li>
	<li>Exchange the scissors for the curved forceps again. Turn the forceps over so that the point is up and away from the animal. With gentle force, push down and dorsal on the rib-cage with the closed, rounded edge of the curved forceps, while pulling down and ventral with the other forceps, spreading the pup out ventral to dorsal. This spreading motion will open the abdominal cavity, exposing the intestines, liver, and other internal organs.</li>
	<li>Using the curved forceps, very gently push the intestines and liver cranially using repeated sweeping motions. This will gradually expose the kidneys, bladder, and most importantly, the internal sex organs.
	<ol>
		<li>Male pups are identified by the presence of striated testes found just below the kidneys on each side. The testes are approximately &#8531;-&#189; the size of the kidneys and occasionally will have dropped to sit just above the bladder.</li>
		<li>Females are most readily identified by a lack of testes, but can also be identified by the small uterus that sits above the bladder, and fallopian tubes that extend up toward the kidneys. Under high-power on the dissection scope, the ovaries can also be identified as small, dark organs at the end of the fallopian tubes immediately below and sometimes underneath the kidneys bilaterally.</li>
	</ol>
	</li>
	<li>Once the sex is determined, discard the body and place the head in an appropriately labeled dish (male or female). Keep the heads on ice in dissection solution.</li>
	<li>Repeat steps 3.5-3.10 until each pup has been sexed. For best results, wipe the dish clean with a Kimwipe between each dissection; fluid makes the pups slippery and difficult to grasp for the dissection.</li>
	<li>Once all embryos have been sexed and the male and female heads have been separated, proceed with the rest of the dissection taking especial care to keep all male and female-derived tissue separate.</li>
	<li>Transfer one brain at a time to a 100 mM dish filled with cold dissection buffer. Under the dissection scope, use the curved forceps to pierce the eyes of the embryo to anchor it for dissection. Using the microsurgical scissors, make a shallow midline incision, splitting the skin and skull down the midline, from base of skull to top of the head just above the nasal passage.</li>
	<li>Gently push aside the skin and skull bilaterally and very gently extract brain by carefully scooping it from the forebrain towards the brainstem with the base of the micro-surgical scissors or with a closed, angled forceps.
	<ol>
		<li>Carefully use the forceps to transfer the extracted brain to a clean, labeled dish filled with cold dissection buffer. Keep the dish on ice.</li>
		<li>Repeat this process until all brains have been harvested.</li>
		<li>This step should yield two plates of whole brains in dissection buffer, one for male and one for female tissue.</li>
	</ol>
	</li>
	<li>Place a clean dish beneath the dissection scope and fill with dissection buffer. Pour just enough solution to keep the brains moist throughout the dissection of the hippocampus; too much will cause the brain to float and make handling difficult. If desired, precoat the dissection dish with sterile wax. The wax provides nonslip surface upon which to work and protects the tips of delicate dissection instruments; scratching the bottom of a noncoated dish can bend or dull the sharp edges of the tools.</li>
	<li>Change the dissection scope to a high-power setting, and adjust the focus accordingly. Carefully transfer one brain at a time to dissection dish. Using the extra-fine tipped forceps, tease apart each brain hemisphere and pinch them off of the brainstem.</li>
	<li>Very carefully, remove the meninges from each hemisphere. (The meninges are a pinkish membrane, rich with blood vessels, and are a common source of glial contamination in a neuron culture.)
	<ol>
		<li>To remove the meninges, delicately pinch, lift, and pull at the membrane, being careful not to pierce the brain tissue beneath.</li>
		<li>Pull the meninges away from the brain, allowing the brain to roll a bit beneath the forceps but keeping it in place so that the membrane can be pulled away.</li>
	</ol>
	</li>
	<li>At this point, you may further dissect the brain to use the tissue of choice. &#160;For example, dissect the hippocampus from each hemisphere. Place one hemisphere with the lateral side down. Observe the large ventricle cavity, which creates a flap covering the hippocampus. Gently tease this flap back until the crescent-shaped hippocampus can be observed. The hippocampus is slightly raised and sits between the mid-brain and cortex.</li>
	<li>Using two pair of sharp, extra-fine forceps, hold the brain hemisphere open with one hand and cut the hippocampus away from the cortex and mid-brain with the other.
	<ol>
		<li>Place the hippocampus in a correctly labeled dish (male or female) filled with dissection buffer and kept on ice.</li>
		<li>Dissect the hippocampus from the remaining hemisphere and repeat this process with each brain.</li>
	</ol>
	</li>
	<li>Transfer the hippocampi into appropriately labeled 15 ml conical tubes.
	<ol>
		<li>Gently wash twice with cold dissection buffer. Use vacuum suction very carefully at this stage. The hippocampi are small and can be irrecoverably lost to the vacuum. Prevent this by allowing the tissue to settle to the bottom of the tube, remove as much liquid as possible from the tube with a pipette, discard the solution, and replace with fresh, clean dissection buffer.</li>
		<li>Repeat step 3.20.1.</li>
	</ol>
	</li>
</ol>
4. Neuronal Dissociation
<ol>
	<li>Transfer the hippocampi to appropriately labeled, prewarmed papain solution. Place in a 37 &#176;C water bath and incubate for 8-10 min, gently inverting the tubes to mix every 2-3 min.</li>
	<li>Remove the papain mix and tissue from the water-bath. Allow the tissue to settle, and then carefully remove as much of the papain solution as possible with a pipette, leaving the tissue at the bottom of the tube. Wash the tissue 3x&#160;with plating media. The serum in the plating media quenches the papain enzymatic digestion. Trypsin inhibitor is needed if serum is not used in the plating media.</li>
	<li>Remove the plating media.
	<ol>
		<li>Add 2 ml&#160;of fresh neurobasal medium and gently dissociate tissue using a 5 ml serological pipette set on the slowest speed.</li>
		<li>Dissociation is performed by gently pipetting up and down 8x. Be careful not to introduce bubbles into solution. For best results, pick the solution and tissue up in the pipette and then release the solution onto the side of the conical tube, allowing the tissue to slide down the side and back to the bottom of the tube.</li>
	</ol>
	</li>
	<li>After the 8th trituration, allow the tissue to settle and collect the cloudy media on top. This media contains disassociated cells.
	<ol>
		<li>Transfer it to a new, labeled tube.</li>
		<li>Add an additional 2 ml to the undisassociated tissue.</li>
		<li>Triturate again another 8x.</li>
		<li>Harvest the cloudy media.</li>
		<li>Repeat this process 3-5x. Some undissociated tissue will remain. While it is possible to triturate until all tissue is dissociated, excessive trituration results in poor neuronal health and is not recommended.</li>
	</ol>
	</li>
	<li>Add extra medium to the conical tubes as necessary to equalize the volumes and weights between male and female tissue. Centrifuge the cells in a spinning bucket centrifuge at 228 x g for 5 min. Carefully remove the supernatant.</li>
	<li>Gently resuspend the cells in 2 ml fresh plating media and filter the solution through a mesh cell strainer into a clean, labeled, 50 ml conical tube.</li>
	<li>Add 50 &#956;l cell suspension to 450 &#956;l of 0.4% Trypan Blue. Incubate the cells for 2 min in Trypan Blue, mix gently, and load 10 &#956;l onto a hemocytometer. Count the cells excluding those that have taken up the Trypan Blue dye; dark blue cells are dead or dying.</li>
</ol>
5. Plating and Culture Maintenance
<ol>
	<li>Calculate the correct volume of cell suspension to add to each well of the tissue culture plate to yield the desired cell density. Neuron cultures are very sensitive to cell density and do best at medium to high density unless cocultured with glia or fed with glial conditioned media. In this protocol, cells are plated at 150,000-200,000 cells/cm2.</li>
	<li>Remove the tissue culture plate prefilled with plating media from the incubator (prepared in step 2 above).
	<ol>
		<li>Gently swirl the cell suspension in the 50 ml conical tube, then pipette the desired volume of cell suspension (calculated above in step 5.1) to the center of each well.</li>
		<li>Repeat, swirling the cell suspension each time before pipetting to ensure equal cell density in each well. Cells settle quickly and cell density can change from well to well if the swirling step is skipped.</li>
	</ol>
	</li>
	<li>Once cells have been added to each well, gently swirl the plate to disperse the cells, and then immediately replace the dish in the incubator. Leave the cells undisturbed for at least 24 hr.</li>
	<li>At least 24 hr&#160;after plating the cells, warm complete neurobasal feeding media to 37 &#176;C. Aspirate the plating media from each well and replace it with feeding media. Aspirate and replace from one well at a time to minimize neuronal stress and improve cell survival.</li>
	<li>Neuron cultures should be fed every 4-7 days by carefully removing half of the medium and adding back an equal volume of fresh medium. Under these conditions, cultures can be maintained for two weeks. With primary neuronal cultures, some cell death is expected. Thus, as the cultures mature, the density will decrease. In this protocol, cell density decreases by approximately 30% by day 10 in vitro.</li>
</ol>

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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=Mechanism+of+the+sex+difference+in+neuronal+ischemic+cell+death.">Mechanism of the sex difference in neuronal ischemic cell death.</a> Neuroscience. 219, 183-191 (2012).</li><li>Verma, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TRPM2+channel+activation+following+in+vitro+ischemia+contributes+to+male+hippocampal+cell+death.">TRPM2 channel activation following in vitro ischemia contributes to male hippocampal cell death.</a> Neurosci. Lett. 530, 41-46 (2012).</li><li>Bai, P., Canto, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+PARP-1+and+PARP-2+enzymes+in+metabolic+regulation+and+disease.">The role of PARP-1 and PARP-2 enzymes in metabolic regulation and disease.</a> Cell Metab. 16, 290-295 (2012).</li><li>Blenn, C., Wyrsch, P., Bader, J., Bollhalder, M., Althaus, F. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Poly(ADP-ribose)glycohydrolase+is+an+upstream+regulator+of+Ca2++fluxes+in+oxidative+cell+death.">Poly(ADP-ribose)glycohydrolase is an upstream regulator of Ca2+ fluxes in oxidative cell death.</a> Cell Mol. Life Sci. 68, 1455-1466 (2011).</li><li>Buelow, B., Song, Y., Scharenberg, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Poly(ADP-ribose)+polymerase+PARP-1+is+required+for+oxidative+stress-induced+TRPM2+activation+in+lymphocytes.">The Poly(ADP-ribose) polymerase PARP-1 is required for oxidative stress-induced TRPM2 activation in lymphocytes.</a> J. Biol. Chem. 283, 24571-24583 (2008).</li><li>Fonfria, E., 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=TRPM2+channel+opening+in+response+to+oxidative+stress+is+dependent+on+activation+of+poly(ADP-ribose)+polymerase.">TRPM2 channel opening in response to oxidative stress is dependent on activation of poly(ADP-ribose) polymerase.</a> Br. J. Pharmacol. 143, 186-192 (2004).</li><li>Miller, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibition+of+TRPM2+function+by+PARP+inhibitors+protects+cells+from+oxidative+stress-induced+death.">Inhibition of TRPM2 function by PARP inhibitors protects cells from oxidative stress-induced death.</a> Br. J. Pharmacol. 143, 515-516 (2004).</li><li>Vagnerova, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Poly+(ADP-ribose)+polymerase-1+initiated+neuronal+cell+death+pathway--do+androgens+matter.">Poly (ADP-ribose) polymerase-1 initiated neuronal cell death pathway--do androgens matter.</a> Neuroscience. , 166-476 (2010).</li><li>Yuan, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sex+differences+in+the+response+to+activation+of+the+poly+(ADP-ribose)+polymerase+pathway+after+experimental+stroke.">Sex differences in the response to activation of the poly (ADP-ribose) polymerase pathway after experimental stroke.</a> Exp. Neurol. 217, 210-218 (2009).</li><li>Berthois, Y., Katzenellenbogen, J. A., Katzenellenbogen, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenol+red+in+tissue+culture+media+is+a+weak+estrogen:+implications+concerning+the+study+of+estrogen-responsive+cells+in+culture.">Phenol red in tissue culture media is a weak estrogen: implications concerning the study of estrogen-responsive cells in culture.</a> Proc. Natl. Acad. Sci. U.S.A. 83, 2496-2500 (1986).</li><li>Dumesic, D. A., Renk, M., Kamel, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estrogenic+effects+of+phenol+red+on+rat+pituitary+cell+responsiveness+to+gonadotropin-releasing+hormone.">Estrogenic effects of phenol red on rat pituitary cell responsiveness to gonadotropin-releasing hormone.</a> Life Sci. 44, 397-406 (1989).</li><li>Ernst, M., Schmid, C., Froesch, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenol+red+mimics+biological+actions+of+estradiol:+enhancement+of+osteoblast+proliferation+in+vitro+and+of+type+I+collagen+gene+expression+in+bone+and+uterus+of+rats+in+vivo.">Phenol red mimics biological actions of estradiol: enhancement of osteoblast proliferation in vitro and of type I collagen gene expression in bone and uterus of rats in vivo.</a> J. Steroid Biochem. 33, 907-914 (1989).</li><li>Hubert, J. F., Vincent, A., Labrie, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estrogenic+activity+of+phenol+red+in+rat+anterior+pituitary+cells+in+culture.">Estrogenic activity of phenol red in rat anterior pituitary cells in culture.</a> Biochem. Biophys. Res. Commun. 141, 885-891 (1986).</li><li>Ortmann, O., Sturm, R., Knuppen, R., Emons, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Weak+estrogenic+activity+of+phenol+red+in+the+pituitary+gonadotroph:+re-evaluation+of+estrogen+and+antiestrogen+effects.">Weak estrogenic activity of phenol red in the pituitary gonadotroph: re-evaluation of estrogen and antiestrogen effects.</a> J. Steroid Biochem. 35, 17-22 (1990).</li><li>Rajendran, K. G., Lopez, T., Parikh, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estrogenic+effect+of+phenol+red+in+MCF-7+cells+is+achieved+through+activation+of+estrogen+receptor+by+interacting+with+a+site+distinct+from+the+steroid+binding+site.">Estrogenic effect of phenol red in MCF-7 cells is achieved through activation of estrogen receptor by interacting with a site distinct from the steroid binding site.</a> Biochem. Biophys. Res. Commun. 142, 724-731 (1987).</li><li>Welshons, W. V., Wolf, M. F., Murphy, C. S., Jordan, V. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estrogenic+activity+of+phenol+red.">Estrogenic activity of phenol red.</a> Mol. Cell Endocrinol. 57, 169-178 (1988).</li><li>Moreno-Cuevas, J. E., Sirbasku, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estrogen+mitogenic+action.+III.+is+phenol+red+a+"red+herring"?.">Estrogen mitogenic action. III. is phenol red a "red herring"?.</a> In Vitro Cell Dev. Biol. Anim. 36, 447-464 (2000).</li></ol>

The authors have nothing to disclose.&#160;

Sex stratified primary disassociated hippocampal neuronal cultures in which male and female hippocampi are dissected, disassociated, and cultured separately present a useful tool for investigating sex-specific cell signaling in neuron cell death. While neuroscientists commonly use primary neuronal cultures, protocols vary between labs depending on their specific experimental aims and interests. Many researchers choose to coculture neurons with glia, either in mixed culture or by plating the neurons on a glass cover slip ...

Mammals are sexually dimorphic with males and females exhibiting different traits and characteristics. Sex differences extend even to the brain and are evident in neuronal susceptibility to injury and disease (reviewed in1-5). For example, the male brain is more susceptible to ischemic neuronal injury from stroke or cardiac arrest followed by resuscitation (reviewed in6). Parkinson&#39;s disease (reviewed in7) and schizophrenia (reviewed in8) are more common in human males than in females, while the female human brain appears more susceptible to Alzheimer&#39;s disease (reviewed in9), and mood disorders such as depression (reviewed in10). Despite observable differences between sexes in susceptibility to neuronal injury and disease, most current treatment paradigms fail to consider sex and are similar in both men and women. Greater success might be achieved if sex differences were accounted for and understood.
In vivo animal models of neuronal injury and disease demonstrate sexual dimorphism similar to humans and are critical to enhancing our understanding of sex differences and how those differences contribute to neuronal survival and function. Evidence has demonstrated, for example, that male rodents are more susceptible than females to ischemic neuronal injury11-15. This difference is thought to be at least partially dependent upon the molecular signaling of testosterone, the primary male sex steroid hormone, and estrogen, the primary female sex steroid hormone. Studies with gonectomized male and female rodents reveals that testosterone and the testosterone metabolite, dihydrotestosterone (DHT) exacerbate and estrogen decreases neuronal injury following cerebral ischemia12-14,16-18. However, sex differences in cerebral ischemia are present in both neonate and adult animals19,20. Neonates experience surges in estrogen or testosterone in utero during development, but have relatively low levels of hormone after birth until puberty (reviewed in21). Thus, the presence or absence of primary sex hormone is not the only determinant of sex-influenced neuronal susceptibility to injury. Indeed, it is likely that sexual dimorphisms are established during development that result in differences in cell signaling and response to ischemia even when hormone levels are low.
Regardless of whether sex differences are dependent on the presence of sex steroid hormone or whether they are due to dimorphisms established during development, it is becoming clear with further study that there are differences in cell signaling between male and females in the brain. In the Herson Lab at the University of Colorado School of Medicine, we have identified one such difference. We have found that the calcium, sodium, and potassium permeable ion channel, TRPM2 (reviewed in22,23), induces neuronal cell death following ischemic insult in vivo in the male but not the female mouse24. We demonstrate that pharmacological inhibition or genetic knockdown of TRPM2 is protective in the male but not the female mouse in our in vivo middle carotid arterial occlusion (MCAO) model of stroke24.
Despite their usefulness, in vivo animal models of ischemic neuronal injury may, at times, be limited for molecular studies of neuronal death, due to the difficulty or inability to specifically target neurons for investigation. For this reason, our lab uses primary disassociated cortical or hippocampal neurons in vitro to complement our in vivo work. Primary disassociated neuron cultures present a useful tool to investigate the cell signaling pathways involved in neuronal injury and disease. Neurons in culture can be maintained in a tightly controlled environment, with or without glial input. In the absence of glia, any responses observed in culture are neuronal. Pharmacological agents can be administered to culture without concerns about absorption across the blood brain barrier, allowing for the inhibition or activation of specific components of a given signal transduction pathway. Molecular biology can be used to overexpress or knockdown various proteins of interest, and electrophysiological measurements can be conducted on individual cells, something that would not be possible in an in vivo model system.
The Herson Lab expands the usefulness of primary disassociated neuronal cultures by first sexing the embryos used to prepare the cultures. Male and female sex-stratified neuron cultures demonstrate clearly that male neurons are more susceptible than female neurons to neuronal cell death following oxygen and glucose deprivation25. Oxygen and glucose deprivation is an accepted in vitro model of neuronal ischemia as ischemia reduces oxygen and glucose availability to the brain. Furthermore, we have demonstrated with electrophysiological experiments in disassociated, sex-stratified neurons that TRPM2 is activated in male but not female neurons24,26. We are currently investigating the molecular sex-specific regulators of TRPM2 activity24,26. Here, we share and discuss our protocol for establishing sex-stratified primary embryonic disassociated hippocampal neuron cultures and present representative data using this method that suggests that the DNA repair enzyme, PARP-1, contributes to ischemic cell death in a sex specific manner similar to TRPM2. PARP-1 is activated by DNA breakage due to oxidative stress (reviewed in27,28). When DNA damage is minimal, PARP-1 activity enhances cell survival, but when damage is excessive, increased PARP-1 activity exacerbates cell death. PARP-1 produces poly-(ADP)ribose, a known activator of the TRPM2 channel28-31, and some evidence suggests that PARP-1 is activated preferentially in the male but not female brain in response to oxidative stress32,33. Thus, we hypothesize that PARP-1 activates TRPM2 in the male but not female, inducing neuronal cell death in response to oxygen and glucose deprivation.

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			<td height="61" style="height:61px;width:227px;">B27 Supplement</td>
			<td style="width:169px;">Invitrogen</td>
			<td style="width:191px;">17504-044</td>
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			<td height="21" style="height:21px;width:227px;">Borax</td>
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			<td height="81" style="height:81px;width:227px;">Bovine Serum Albumin</td>
			<td style="width:169px;">Sigma</td>
			<td style="width:191px;">A4503</td>
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			<td height="61" style="height:61px;width:227px;">Fetal Bovine Serum</td>
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			<td height="41" style="height:41px;width:227px;">GlutaMAX</td>
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			<td style="width:191px;">12348-017</td>
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			<td height="21" style="height:21px;width:227px;">Penicillin/Streptomycin</td>
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			<td style="width:191px;">15070-063</td>
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			<td height="41" style="height:41px;width:227px;">Poly-D-lysine, MW 70,000-150,000</td>
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			<td height="21" style="height:21px;width:227px;">Trypan Blue 0.4%</td>
			<td style="width:169px;">Invitrogen</td>
			<td style="width:191px;">15250-061</td>
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			<td style="width:169px;">Fine Science Tools</td>
			<td style="width:191px;">11251-35</td>
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		<tr height="41">
			<td height="41" style="height:41px;width:227px;">Dumont #7 &#8211; Fine Curved Forceps</td>
			<td style="width:169px;">Fine Science Tools</td>
			<td style="width:191px;">11274-20</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:227px;">Serrated (London) Forceps</td>
			<td style="width:169px;">Fine Science Tools</td>
			<td style="width:191px;">11080-02</td>
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		<tr height="41">
			<td height="41" style="height:41px;width:227px;">Extra Fine Graefe Forceps 1 x 2 teeth</td>
			<td style="width:169px;">Fine Science Tools</td>
			<td style="width:191px;">11153-10</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:227px;">Moria Spring Scissors</td>
			<td style="width:169px;">Fine Science Tools</td>
			<td style="width:191px;">15396-00</td>
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			<td height="21" style="height:21px;width:227px;">Medium Operating Scissors</td>
			<td style="width:169px;">Roboz Surgical</td>
			<td style="width:191px;">RS-6812</td>
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			<td height="41" style="height:41px;width:227px;">Extra Fine Microdissecting Scissors</td>
			<td style="width:169px;">Roboz Surgical</td>
			<td style="width:191px;">RS-5880</td>
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			<td height="21" style="height:21px;width:227px;">24-well Tissue Culture Plates</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:191px;">87721</td>
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			<td height="41" style="height:41px;width:227px;">Cell Strainer, 40 &#181;M Nylon Mesh</td>
			<td style="width:169px;">BD Biosciences</td>
			<td style="width:191px;">352340</td>
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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.

Proper sexing of embryonic mice is a critical step in the preparation of sex-stratified primary neuron cultures. To ensure the male and female hippocampi were cultured separately from correctly sexed pups, embryos were separated by sex and their cortex and hippocampus were harvested. Genomic DNA was isolated from the harvested tissue of each embryo according to standard procedures, being careful to keep sex separate. Polymerase Chain Reaction (PCR) was performed using primers for the male-specific gene, SRY and the unive...

Watch this Scientific Journal Video about Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways at JoVE.com

Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways

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 ...

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Neuroscience

11.9K Views.  University of Colorado School of Medicine. 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.

Video: Sex Stratified Neuronal Cultures to Study Ischemic Cell Death Pathways

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

The mouse is an excellent model organism to study mammalian brain development due to the abundance of molecular and genetic data. However, the developing mouse brain is not suitable for easy manipulation and imaging in vivo since the mouse embryo is inaccessible and opaque. Organotypic slice cultures of embryonic brains are therefore widely used to study murine brain development in vitro. Ex-vivo manipulation or the use of transgenic mice allows the modification of gene expression so that subpopulations of neuronal or glial cells can be labeled with fluorescent proteins. The behavior of labeled cells can then be observed using time-lapse imaging. Time-lapse imaging has been particularly successful for studying cell behaviors that underlie the development of the cerebral cortex at late embryonic stages 1-2. Embryonic organotypic slice culture systems in brain regions outside of the forebrain are less well established. Therefore, the wealth of time-lapse imaging data describing neuronal cell migration is restricted to the forebrain 3,4. It is still not known, whether the principles discovered for the dorsal brain hold true for ventral brain areas. In the ventral brain, neurons are organized in neuronal clusters rather than layers and they often have to undergo complicated migratory trajectories to reach their final position. The ventral midbrain is not only a good model system for ventral brain development, but also contains neuronal populations such as dopaminergic neurons that are relevant in disease processes. While the function and degeneration of dopaminergic neurons has been investigated in great detail in the adult and ageing brain, little is known about the behavior of these neurons during their differentiation and migration phase 5. We describe here the generation of slice cultures from the embryonic day (E) 12.5 mouse ventral midbrain. These slice cultures are potentially suitable for monitoring dopaminergic neuron development over several days in vitro. We highlight the critical steps in generating brain slices at these early stages of embryonic development and discuss the conditions necessary for maintaining normal development of dopaminergic neurons in vitro. We also present results from time lapse imaging experiments. In these experiments, ventral midbrain precursors (including dopaminergic precursors) and their descendants were labeled in a mosaic manner using a Cre/loxP based inducible fate mapping system 6.

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

A method to generate organotypic slices from the E12.5 murine embryonic midbrain is described. The organotypic slice cultures can be used to observe the behavior of dopaminergic neurons or other ventral midbrain neurons.

The  mouse is an excellent model organism to study mammalian brain development due  to the abundance of molecular and genetic data. However, the ...

Preparation of Neuronal Co-cultures with Single Cell Precision

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms can be used to investigate a variety of questions, spanning developmental and functional neurobiology to infection and disease propagation. However, conventional systems require significant cellular inputs (many thousands per compartment), inadequate for studying low abundance cells, such as primary dopaminergic substantia nigra, spiral ganglia, and Drosophilia melanogaster neurons, and impractical for high throughput experimentation. The dense cultures are also highly locally entangled, with few outgrowths (&#60;10%) interconnecting the two cultures. In this paper straightforward microfluidic and patterning protocols are described which address these challenges: (i) a microfluidic single neuron arraying method, and (ii) a water masking method for plasma patterning biomaterial coatings to register neurons and promote outgrowth between compartments. Minimalistic neuronal co-cultures were prepared with high-level (&#62;85%) intercompartment connectivity and can be used for high throughput neurobiology experiments with single cell precision.

Protocols for single neuron microfluidic arraying and water masking for the in-chip plasma patterning of biomaterial coatings are described. Highly interconnected co-cultures can be prepared using minimal cell inputs.

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms ...

Real-Time Impedance-based Cell Analyzer as a Tool to Delineate Molecular Pathways Involved in Neurotoxicity and Neuroprotection in a Neuronal Cell Line

Many brain-related disorders have neuronal cell death involved in their pathophysiology. Improved in vitro models to study neuroprotective or neurotoxic effects of drugs and downstream pathways involved would help gain insight into the molecular mechanisms of neuroprotection/neurotoxicity and could potentially facilitate drug development. However, many existing in vitro toxicity assays have major limitations &#8211; most assess neurotoxicity and neuroprotection at a single time point, not allowing to observe the time-course and kinetics of the effect. Furthermore, the opportunity to collect information about downstream signaling pathways involved in neuroprotection in real-time would be of great importance. In the current protocol we describe the use of a real-time impedance-based cell analyzer to determine neuroprotective effects of serotonin 2A (5-HT2A) receptor agonists in a neuronal cell line under label-free and real-time conditions using impedance measurements. Furthermore, we demonstrate that inhibitors of second messenger pathways can be used to delineate downstream molecules involved in the neuroprotective effect. We also describe the utility of this technique to determine whether an effect on cell proliferation contributes to an observed neuroprotective effect. The system utilizes special microelectronic plates referred to as E-Plates which contain alternating gold microelectrode arrays on the bottom surface of the wells, serving as cell sensors. The impedance readout is modified by the number of adherent cells, cell viability, morphology, and adhesion. A dimensionless parameter called Cell Index is derived from the electrical impedance measurements and is used to represent the cell status. Overall, the real-time impedance-based cell analyzer allows for real-time, label-free assessment of neuroprotection and neurotoxicity, and the evaluation of second messenger pathways involvement, contributing to more detailed and high-throughput assessment of potential neuroprotective compounds in vitro, for selecting therapeutic candidates.

Improved in vitro neurotoxicity assays would aid the identification of new neuroprotective compounds. The utility of a real-time impedance-based cell analyzer to determine cytotoxicity and cytoprotection in neuronal cell lines and to delineate the involvement of second messenger pathways, thus gaining insight in the mechanism of neuroprotection is presented.

Many brain-related disorders have neuronal cell death involved in their pathophysiology. Improved in vitro models to study neuroprotective or neurotoxic ...

Organotypic Slice Cultures to Study Oligodendrocyte Dynamics and Myelination

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During embryonic and postnatal development they actively proliferate and generate myelinating oligodendrocytes. These cells have commonly been studied in primary dissociated cultures, neuron cocultures, and in fixed tissue. Using newly available transgenic mouse lines slice culture systems can be used to investigate proliferation and differentiation of oligodendrocyte lineage cells in both gray and white matter regions of the forebrain and cerebellum. Slice cultures are prepared from early postnatal mice and are kept in culture for up to 1 month. These slices can be imaged multiple times over the culture period to investigate cellular behavior and interactions. This method allows visualization of NG2 cell division and the steps leading to oligodendrocyte differentiation while enabling detailed analysis of region-dependent NG2 cell and oligodendrocyte functional heterogeneity. This is a powerful technique that can be used to investigate the intrinsic and extrinsic signals influencing these cells over time in a cellular environment that closely resembles that found in vivo.

A technique to study NG2 cells and oligodendrocytes using a slice culture system of the forebrain and cerebellum is described. This method allows examination of the dynamics of proliferation and differentiation of cells within the oligodendrocyte lineage where the extracellular environment can be easily manipulated while maintaining tissue cytoarchitecture.

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During ...

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, ...

Organotypic Hippocampal Slice Cultures As a Model to Study Neuroprotection and Invasiveness of Tumor Cells

In organotypic hippocampal slice cultures (OHSC), the morphological and functional characteristics of both neurons and glial cells are well preserved. This model is suitable for addressing different research questions that involve studies on neuroprotection, electrophysiological experiments on neurons, neuronal networks or tumor invasion. The hippocampal architecture and neuronal activity in multisynaptic circuits are well conserved in OHSC, even though the slicing procedure itself initially lesions and leads to formation of a glial scar. The scar formation alters presumably the mechanical properties and diffusive behavior of small molecules, etc. Slices allow the monitoring of time dependent processes after brain injury without animal surgery, and studies on interactions between various brain-derived cell types, namely astrocytes, microglia and neurons under both physiological and pathological conditions. An ambivalent aspect of this model is the absence of blood flow and immune blood cells. During the progression of the neuronal injury, migrating immune cells from the blood play an important role. As those cells are missing in slices, the intrinsic processes in the culture may be observed without external interference. Moreover, in OHSC the composition of the medium-external environment is precisely controlled. A further advantage of this method is the lower number of sacrificed animals compared to standard preparations. Several OHSC can be obtained from one animal making simultaneous studies with multiple treatments in one animal possible. For these reasons, OHSC are well suited to analyze the effects of new protective therapeutics after tissue damage or during tumor invasion. 
The protocol presented here describes a preparation method of OHSC that allows generating highly reproducible, well preserved slices that can be used for a variety of experimental research, like neuroprotection or tumor invasion studies.

Organotypic hippocampal slice cultures (OHSC) represent an in vitro model that simulates the in vivo situation very well. Here we describe a vibratome-based improved slicing protocol to obtain high quality slices for use in assessing the neuroprotective potential of novel substances or the biological behavior of tumor cells.

In organotypic hippocampal slice cultures (OHSC), the morphological and functional characteristics of both neurons and glial cells are well preserved. ...

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 ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

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.

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 ...

Hippocampal Neuronal Cultures to Detect and Study New Pathogenic Antibodies Involved in Autoimmune Encephalitis

Over the last 15 years, a new category of antibody-mediated diseases of the central nervous system (CNS) has been characterized and is now defined as &#34;autoimmune encephalitis&#34; (AE). There are currently 17 known AE syndromes, and all are associated with antibodies against the neuronal cell surface or synaptic proteins. The clinical syndromes are complex and vary according to the type of associated antibody. The best-known of these diseases is anti-N-methyl D-aspartate receptor (NMDAR) encephalitis, which is a prominent neuropsychiatric disorder associated with severe memory and behavioral impairments. The associated antibodies react with the GluN1 subunit of the NMDAR at the N-terminal domain. The approach most frequently used for the discovery and characterization of AE antibodies includes the culture of dissociated, fetal, rodent hippocampal neurons. During the process of antibody characterization, live neurons in culture are exposed to patients&#39; serum or CSF, and the detection of reactivity indicates that the serum or CSF samples of the patient contain antibodies against neuronal surface antigens. Hippocampal cultures can also be used to determine whether the antibodies in patients are potentially pathogenic by examining if they cause structural or functional alterations of the neurons. The level of success of these studies depends on the quality of the cultures and on the protocols used to obtain and detect the reactivity of patient samples. This article provides an optimized protocol for primary cell culture of fetal rat hippocampal neurons combined with immunostaining to determine the presence of antibodies in the serum or CSF of patients. An example of how to examine the potential pathogenic effects of NMDAR antibodies using cultured neurons and calcium imaging is also presented.

Autoimmune encephalitis is a new category of antibody-mediated diseases of the central nervous system. Hippocampal neurons can be used to discover and characterize these antibodies. This article provides a protocol for primary cell culture and immunostaining to determine autoantibodies in the serum and cerebrospinal fluid of patients.

Over the last 15 years, a new category of antibody-mediated diseases of the central nervous system (CNS) has been characterized and is now defined as ...