This work was supported by grants from Canadian Institutes of Health Research MOP-81158.

1. Extraction of Target Protein from Cultured Primary Hippocampal Neurons
<ol>
 <li>Before harvesting endogenous protein from primary hippocampal cells, prepare a Lysis Buffer (LB; Table 1) with protease inhibitors. To 10 ml of lysis buffer, add 0.1 ml phenylmethanesulfonyl fluoride solution (PMSF) to a final concentration of 10mM, and add 1 protease inhibitor cocktail tablet.</li>
 <li>Prepare N-ethylmaleimide (NEM) solution (Table 1) from lyophilized powder. Gently dissolve in 100% EtOH at room temperature (RT). Always prepare NEM solution fresh for the day of the experiment.</li>
 <li>Next combine the lysis buffer and NEM. Add 0.5 ml of 2M NEM solution to a fresh 50 ml conical tube. Add 20 ml LB over top, and quickly place LB +NEM (final concentration of 50mM) on a rocking platform at 4 &deg;C to fully mix for 5 min. Always add NEM/EtOH solution first and LB second, so as to properly mix the two.</li>
 <li>Remove cultured neurons from incubator and place on ice*. Gently remove cellular media, and gently wash twice with cold phosphate buffered saline (PBS) 1x. Add 300 ml of LB + 50mM NEM to the first well of hippocampal neurons, and scrape cells using a disposable cell lifter. Collect the cell lysate, then discharge the lysate into the next well of that group. Scrape cells within the second well using the cell lifter, collect, and repeat using a third well, if needed to further concentrate the lysate.</li>
 <li>Collect cell lysate into a pre-cooled (on ice), 1.5 ml microcentrifuge tube. Repeat step 1.4 using 100 &mu;l of LB + 50mM NEM to collect any remaining cell lysate. Pass total cell lysate through a 26&frac12; gauge syringe 5-6 times to aid in mechanical lysis, being careful to not blow bubbles into the solution. One can instead sonicate for 15 sec on ice, if equipment is available. Nutate cell lysate for &ge;30 min at 4 &deg;C.</li>
 <li>Clarify the cell lysate by centrifugation at 16,000 x g/ 30 min, in order to pellet un-lysed cells and detergent-insoluble material.</li>
 <li>Collect cleared cell lysate (supernatant) in a new pre-cooled 1.5 ml tube on ice. Repeat harvest protocol for all groups of cells. Measure protein concentration in all lysate samples using BCA assay, and normalize the volume of all lysate samples from groups of cells to have exactly equal total protein, with a recommended minimum of 500 &mu;g. Top up all samples with LB + 50mM NEM to 500 &mu;l total volume.</li>
 <li>Add 1-5 &mu;g of primary antibody directed against the target protein to each of lysate sample. Nutate the antibody-lysate mixed samples overnight at 4 &deg;C.</li>
</ol>
* Primary rat hippocampal neurons should be cultured at a density of approximately 250,000 cells/well in a 6-well dish, in a serum-free media containing a B27 neuronal supplement, as previously described5, and grown for the desired length of time, typically 14-16 days-in-vitro (DIV) to achieve maturity. A minimum of 500 &mu;g of total protein is recommended to successfully immunoprecipitate and biotinylate a target neuronal protein, which typically requires 2-3 wells of a 6-well dish.
2. Precipitation of Antibody-bound Target Protein
<ol>
 <li>Before precipitating and immobilizing a target protein, prepare a 50% slurry of protein A, or protein G-coated sepharose beads. Specifically, add &ge;60 &mu;l of sepharose beads per sample to 1.5 ml tubes, ensuring that all samples have equal amounts of beads. Magnetic beads are also suitable if the equipment is available.</li>
 <li>Add an equal volume of 50% slurry to each antibody-lysate sample, and nutate for &ge;1 hr at 4 &deg;C.</li>
</ol>
3. Acyl-Biotin Exchange: Hydroxylamine (HAM) Cleavage
<ol>
 <li>While performing step 2.2, prepare a number of tubes with lysis buffer (LB) of different pHs. The pH is very important for these steps and should always be adjusted using a pH meter. Prepare 2 ml of LB pH 7.2 per sample, and 0.5 ml of Stringent Buffer per sample (Table 1). Also prepare 0.5 ml LB + 10mM NEM per sample, as in steps 1.1-1.3. Add PMSF and protease inhibitor tablets to all lysis buffers, as in step 1.1.</li>
 <li>
 <ol class="lalpha">
 <li>Hydroxylamine (HAM) is a powerful reducing agent, whose cleavage of palmitate from cysteines is required for biotinylation (Figure 1), and the omission of the HAM cleavage serves as a negative control. Split each sample of beads into two samples, one omitting the HAM cleavage step (-HAM), and one including the HAM step (+HAM). To normalize for protein degradation caused by HAM treatment, one should split each sample into thirds, with 1/3 of the beads used for the -HAM control, and the remaining 2/3 used for the +HAM treatment.</li>
 <li>Prepare additional 1.5 ml tubes on ice, labeled as the -HAM control for each sample. Following step 2.2, gently centrifuge all samples' beads at 0.5 x g/ 1 min at 4 &deg;C (centrifuge at this speed, duration, and temperature for all remaining steps unless otherwise stated), place the tubes on ice, remove the supernatant, and re-suspend the beads in 600 &mu;l of LB + 10mM NEM.</li>
 </ol>
 </li>
 <li>After re-suspending the samples' beads in lysis buffer + NEM 10mM, immediately collect 200 &mu;l of the sample's lysate-bead slurry, and discharge into that sample's -HAM tube on ice. Add an additional 300 &mu;l of LB + 10mM NEM to the -HAM sample, and an additional 100 &mu;l to the +HAM sample for a total of 500 &mu;l in each sample. Incubate the tubes for 10 min on ice.</li>
 <li>Re-suspend (wash) the -HAM and +HAM samples gently with 0.5 ml stringent buffer, per sample. Perform this step quickly, as a longer SDS incubation will result in the removal of bound antibody from the beads.</li>
 <li>Gently wash all samples three times in 0.5 ml/sample of LB pH 7.2, and spin down between washes as in step 3.2b.</li>
 <li>While performing step 3.5, prepare a HAM buffer (Table 1). Add appropriate volume of hydroxylamine solution to a new tube, to achieve a final concentration of 1M in a volume of 0.5 ml LB pH 7.2, per +HAM sample. Always prepare the HAM buffer fresh for the day of the experiment. Add 0.5 ml/sample of LB pH 7.2 to all -HAM samples, and 0.5 ml/ sample of HAM buffer to all +HAM samples.</li>
 <li>Nutate all samples for 1 hr at room temperature (RT). Do not exceed 1 hr.</li>
</ol>
4. Acyl-Biotin Exchange: Biotin-BMCC Labeling
<ol>
 <li>While performing step 3.6, prepare 2 ml of LB pH 6.2 per sample (Table 1), as in step 3.1. Place the LB pH 6.2 on ice until needed. Also prepare a stock solution of Biotin-BMCC (Table 1) immediately before use. Weigh out exactly 2.1 mg of Biotin-BMCC, collect in a 1.5 ml tube, and gently dissolve in 0.5 ml Dimethyl sulfoxide (DMSO) by placing on a rocking platform at RT, to achieve 8mM concentration (do not use a pipette to break up the powder).</li>
 <li>Once step 3.6 is completed, gently wash the beads once in pH 6.2 lysis buffer, to remove residual HAM buffer. Remove the supernatant and place the samples on ice.</li>
 <li>Biotin-BMCC is a maliemide conjugated biotin molecule that has a high affinity for free thiol groups of cysteines (Figure 1). While performing step 4.2, prepare 0.5 ml of Biotin-BMCC Buffer (Table 1) per sample, at working concentration between 0.5 &mu;M to 5 &mu;M. Concentrations in excess of 5 &mu;M will likely saturate the target protein6, and typically should not be exceeded, however this may differ depending on the desired target protein. Add 0.5 ml of Biotin-BMCC buffer to each sample, and nutate for exactly 1 hr at 4 &deg;C. Do not exceed 1 hr.</li>
</ol>
5. Elution of Biotin-conjugated Target Protein and SDS-PAGE
<ol>
 <li>Once step 4.3 is completed, gently wash all samples once in LB pH 6.2. Keep all samples on ice in between washes. Remove 2x SDS Sample Buffer with no reducing agents (Table 1) from -20 &deg;C storage and slowly thaw on ice.</li>
 <li>Gently wash samples three times in LB pH 7.5 + PMSF/Inhibitors, and keep samples on ice in between washes.</li>
 <li>Remove the supernatant of the third wash, leaving the pelleted beads in a slurry with remaining buffer in the bottom of the tube. Dip a pipette with a small-diameter tip (a gel loading tip or P2 pipette tip are sufficient) into the very bottom of the tube through the slurry, and quickly collect any remaining buffer, being careful not to take up any beads.</li>
 <li>Supplement the 2x SDS Sample Buffer with DL-Dithiothreitol (DTT) to achieve a final concentration of 5mM. Add 40-50 &mu;l of 2x Sample Buffer + 5mM DTT to each sample. If necessary, vortex the beads to completely mix with the sample buffer, and immediately centrifuge at high speed to collect all beads in a pellet, submerged in sample buffer.</li>
 <li>Boil all samples for 10 min at 75-80 &deg;C. Let samples cool to RT for &ge;10 min on bench top, and centrifuge at 16,000 x g / 3 min to completely pellet the beads.</li>
 <li>Add complete volume of eluted protein (supernatant) from each sample to a single lane within a polyacrylamide gel suitable for western blotting, using SDS-PAGE7. Organize all samples so that the -HAM control eluent and +HAM eluent for a single sample are run immediately adjacent to one another during SDS-PAGE (Fig. 2). Following SDS-PAGE, transfer to a PVDF or nitrocellulose membrane.</li>
</ol>
6. Western Blotting and Detection of Palmitoylation of Target Protein
<ol>
 <li>Once step 5.6 is completed, wash the membrane twice in Tris-buffered Saline (TBS 1x) for 10 min on a rocking platform at RT.</li>
 <li>Prepare a Blocking Buffer to block non-specific labeling by weighing out Bovine Serum Albumin (BSA), to achieve 3% weight in a volume of TBS 1x + Tween-20 0.05% (TBS-T) that is sufficient to fully submerge the membrane. For streptavidin blotting, always block with BSA and not powdered milk. Block the membrane for 1 hr at RT on a rocking platform.</li>
 <li>Prepare an antibody solution of TBS-T + 0.3% BSA. Add Streptavidin-HRP antibody reconstituted at 1 mg/ml in distilled water, diluted at 1:5,000-1:10,000. Once step 6.2 is completed, wash the membrane once in TBS-T for 10 min, then incubate the membrane in Streptavidin antibody solution on a rocking platform for either for 1 hr at RT, or overnight at 4 &deg;C. </li>
 <li>Wash the membrane three times in TBS-T for 10 min on a rocking platform. In order to detect the palmitoylation signal, expose the HRP luminescence using a chemiluminescent substrate kit.</li>
 <li>In order to normalize palmitoylation levels to the amount of the protein of interest that was immunoprecipitated, strip the membrane using western blot stripping buffer for 5-10 min on a rocking platform at RT. Repeat steps 6.1 - 6.4 using the primary antibody against the protein of interest that was originally used for the immunoprecipitation.</li>
 <li>Quantify palmitoylation of the protein of interest using image analysis software, and normalize palmitoylation levels to the amount of immunoprecipitated protein.</li>
</ol>

<ol>
	<li>Fukata, Y., Fukata, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+palmitoylation+in+neuronal+development+and+synaptic+plasticity.">Protein palmitoylation in neuronal development and synaptic plasticity.</a> Nat. Rev. Neurosci. 11, 161-175 (2010).</li><li>Kang, R., 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=Neural+palmitoyl-proteomics+reveals+dynamic+synaptic+palmitoylation.">Neural palmitoyl-proteomics reveals dynamic synaptic palmitoylation.</a> Nature. 456, 904-909 (2008).</li><li>Drisdel, R. C., Green, W. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Labeling+and+quantifying+sites+of+protein+palmitoylation.">Labeling and quantifying sites of protein palmitoylation.</a> Biotechniques. 36, 276-285 (2004).</li><li>Wan, J., Roth, A. F., Bailey, A. O., Davis, N. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Palmitoylated+proteins:+purification+and+identification.">Palmitoylated proteins: purification and identification.</a> Nat. Protoc. 2, 1573-1584 (2007).</li><li>Xie, C., Markesbery, W. R., Lovell, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+of+hippocampal+and+cortical+neurons+in+a+mixture+of+MEM++and+B27-supplemented+neurobasal+medium.">Survival of hippocampal and cortical neurons in a mixture of MEM+ and B27-supplemented neurobasal medium.</a> Free Radic. Biol. Med. 28, 665-672 (2000).</li><li>Drisdel, R. C., Alexander, J. K., Sayeed, A., Green, W. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assays+of+protein+palmitoylation.">Assays of protein palmitoylation.</a> Methods. 40, 127-134 (2006).</li><li>Eslami, A., Lujan, J. W. e. s. t. e. 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=Blotting:+Sample+Preparation+to+Detection.">Blotting: Sample Preparation to Detection.</a> J. Vis. Exp. (44), e2359 (2010).</li><li>Huang, 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=Neuronal+palmitoyl+acyl+transferases+exhibit+distinct+substrate+specificity.">Neuronal palmitoyl acyl transferases exhibit distinct substrate specificity.</a> FASEB J. 23, 2605-2615 (2009).</li><li>Noritake, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mobile+DHHC+palmitoylating+enzyme+mediates+activity-sensitive+synaptic+targeting+of+PSD-95.">Mobile DHHC palmitoylating enzyme mediates activity-sensitive synaptic targeting of PSD-95.</a> J. Cell Biol. 186, 147-160 (2009).</li><li>Thomas, G. M., Hayashi, T., Chiu, S. L., Chen, C. M., Huganir, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Palmitoylation+by+DHHC5/8+Targets+GRIP1+to+Dendritic+Endosomes+to+Regulate+AMPA-R+Trafficking.">Palmitoylation by DHHC5/8 Targets GRIP1 to Dendritic Endosomes to Regulate AMPA-R Trafficking.</a> Neuron. 73, 482-496 (2012).</li><li>Hayashi, T., Thomas, G. M., Huganir, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+palmitoylation+of+NR2+subunits+regulates+NMDA+receptor+trafficking.">Dual palmitoylation of NR2 subunits regulates NMDA receptor trafficking.</a> Neuron. 64, 213-226 (2009).</li><li>Ho, G. P., 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=S-Nitrosylation+and+S-Palmitoylation+Reciprocally+Regulate+Synaptic+Targeting+of+PSD-95.">S-Nitrosylation and S-Palmitoylation Reciprocally Regulate Synaptic Targeting of PSD-95.</a> Neuron. 71, 131-141 (2011).</li><li>Iwanaga, T., Tsutsumi, R., Noritake, J., Fukata, Y., Fukata, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+protein+palmitoylation+in+cellular+signaling.">Dynamic protein palmitoylation in cellular signaling.</a> Prog. Lipid Res. 48, 117-127 (2009).</li></ol>

No conflicts of interest declared.

The IP-ABE assay presented here is a simple and highly sensitive method for detecting palmitoylated proteins in cultured primary hippocampal neurons, using mostly standard biochemical techniques. This makes the assay easily adaptable for labs already equipped with western blotting materials. The IP-ABE assay combines a standard immunoprecipitation protocol to isolate and immobilize one's target protein, followed by previously described ABE chemistry2-4 to rapidly detect levels of palmitate on a substrate protein. The ABE technique has a wide range of applications for examining palmitoylated proteins in various tissues and cell lines, including large-scale palmitoyl-proteomic screening of global palmitoylation profiles, and rapid detection of palmitoylation levels of a single target protein.
Previous descriptions of ABE chemistry have utilized different methods cell lysis and detergent extraction of neuronal proteins2, 9, free thiol-blockade10, 11, biotinylation, and purification of the target protein4, depending on the application of the experiment. The addition of palmitate to neuronal proteins results in their targeting toward cellular membranes, and detection of the palmitoylated fraction of a target protein will require its extraction from these membranes. Previously described ABE methodologies have utilized a variety of ionic9 and non-ionic detergents8 to extract desired target proteins, and the use of a particular detergent depends on the target protein and its known membrane trafficking. Here, we utilize the non-denaturing and non-ionic detergent IGEPAL CA-630 to extract palmitoylated proteins, which we have validated for extraction and detection of the palmitoylated neuronal protein &delta;-catenin (Figure 2). The structure of IGEPAL CA-630 includes a bulky and rigid non-polar head group that does not disrupt native protein conformation or interactions, but which allows its association with the hydrophobic regions of membrane-associated proteins, thereby conferring miscibility to them and permitting their extraction. Many neuronal proteins localized to the synaptic membrane or post-synaptic density of synapses should be extracted using IGEPAL CA-630, however some may not completely solubilize, and may require use of a different non-denaturing detergent like Triton X-100, which has been previously utilized for ABE chemistry2, 8.
Several descriptions of ABE chemistry have also utilized a different thiol-reactive biotinylation reagent from the molecule we describe here, called Biotin-HPDP (Thermo Scientific), which also necessitates a different means of protein purification4. Biotin-HPDP is another thiol-reactive, maliemide-conjugated biotin molecule that contains a disulfide bond in the maliemide linker arm, structurally differentiating it from Biotin-BMCC, which does not contain this disulfide bond. Following thiol-biotinylation of a protein's free or palmitoylated cysteine by Biotin-HPDP, the resulting disulfide bond between the target protein and the biotin group can be cleaved by reducing reagents to release the biotin group and regenerate the protein in its unmodified form, making biotinylation by Biotin-HPDP transient and reversible. Therefore, use of this biotinylation reagent requires purification of a target protein by immobilized avidin reagents, such as streptavidin-coated beads. However, thiol-biotinylation of a target protein by Biotin-BMCC, as we describe here, is stable and relatively permanent, and requires purification of the target protein by an antibody directed against the target, and immobilized on sepharose beads. Both ABE techniques have been previously utilized for large-scale screens, and detection of palmitoylation of single proteins2, 8-10, 12, and the IP-ABE assay we describe here is optimal for rapid detection of palmitoylation of single neuronal target proteins.
An overwhelming majority of cellular palmitoylation involves the reversible addition of palmitate to the thiol group of cysteines (termed S-palmitoylation, which we generalize as 'palmitoylation' in this protocol), however a very limited group of proteins are subjected to irreversible palmitoylation at glycine and cysteine residues through the formation of stable amide bonds, termed N-palmitoylation13. One limitation of ABE chemistry is its inability to detect N-palmitoylation owing to the stability of the amide bond, and so screening of a novel target protein for palmitoylation should be confirmed using 3H-palmitate metabolic labeling1.
As previously mentioned, the IP-ABE assay can be easily adapted for examining palmitoylation in protein extracts from multiple sources, including transfected heterologous cell lines, primary tissue, and primary neuronal cultures from multiple brain regions. The IP-ABE assay has been utilized for examining palmitoylation sufficiency of mutated cysteine-to-serine proteins to identify the location of a palmitoylated cysteine along a target protein8, for examining dynamic palmitate turnover on synaptic proteins in cultured neurons under basal conditions10, and changes in palmitoylation levels of neuronal proteins following induction of neuronal activity9. The sensitivity and adaptability of the IP-ABE assay make it ideally suited for examining palmitoylation profiles of neuronal proteins, and optimal for detecting dynamic changes in palmitoylation.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>IGEPAL CA-630</td>
 <td>Sigma</td>
 <td>I8896</td>
 <td></td>
 </tr>
 <tr>
 <td>PMSF</td>
 <td>Fluka </td>
 <td>93482</td>
 <td></td>
 </tr>
 <tr>
 <td>Protease Inhibitor Cocktail</td>
 <td>Roche Complete Mini</td>
 <td>11 836 153 001</td>
 <td></td>
 </tr>
 <tr>
 <td>NEM</td>
 <td>Sigma</td>
 <td>E3876</td>
 <td></td>
 </tr>
 <tr>
 <td>HAM solution </td>
 <td>Sigma</td>
 <td>46780-4</td>
 <td></td>
 </tr>
 <tr>
 <td>BCA Protein Assay Kit</td>
 <td>Thermo Scientific </td>
 <td>23225</td>
 <td>Ensure compatibility with chosen detergent</td>
 </tr>
 <tr>
 <td>Protein G/A-coated sepharose beads </td>
 <td>GE Healthcare</td>
 <td>17-6002-35</td>
 <td></td>
 </tr>
 <tr>
 <td>Biotin-BMCC</td>
 <td>Thermo Scientific</td>
 <td>21900</td>
 <td></td>
 </tr>
 <tr>
 <td>Streptavidin-HRP</td>
 <td>Thermo Scientific</td>
 <td>21126</td>
 <td>Reconstitute at 1 mg/ml in water</td>
 </tr>
 <tr>
 <td>Western Blot Stripping Buffer</td>
 <td>Thermo Scientific</td>
 <td>21059</td>
 <td></td>
 </tr>
 </tbody>
</table>

detection of protein palmitoylation in cultured hippocampal neurons by immunoprecipitation and acyl-biotin exchange (abe)

Palmitoylation is a post-translational lipid modification involving the attachment of a 16-carbon saturated fatty acid, palmitate, to cysteine residues of substrate proteins through a labile thioester bond [reviewed in1]. Palmitoylation of a substrate protein increases its hydrophobicity, and typically facilitates its trafficking toward cellular membranes. Recent studies have shown palmitoylation to be one of the most common lipid modifications in neurons1, 2, suggesting that palmitate turnover is an important mechanism by which these cells regulate the targeting and trafficking of proteins. The identification and detection of palmitoylated substrates can therefore better our understanding of protein trafficking in neurons.
Detection of protein palmitoylation in the past has been technically hindered due to the lack of a consensus sequence among substrate proteins, and the reliance on metabolic labeling of palmitoyl-proteins with 3H-palmitate, a time-consuming biochemical assay with low sensitivity. Development of the Acyl-Biotin Exchange (ABE) assay enables more rapid and high sensitivity detection of palmitoylated proteins2-4, and is optimal for measuring the dynamic turnover of palmitate on neuronal proteins. The ABE assay is comprised of three biochemical steps (Figure 1): 1) irreversible blockade of unmodified cysteine thiol groups using N-ethylmaliemide (NEM), 2) specific cleavage and unmasking of the palmitoylated cysteine's thiol group by hydroxylamine (HAM), and 3) selective labeling of the palmitoylated cysteine using a thiol-reactive biotinylation reagent, biotin-BMCC. Purification of the thiol-biotinylated proteins following the ABE steps has differed, depending on the overall goal of the experiment.
Here, we describe a method to purify a palmitoylated protein of interest in primary hippocampal neurons by an initial immunoprecipitation (IP) step using an antibody directed against the protein, followed by the ABE assay and western blotting to directly measure palmitoylation levels of that protein, which is termed the IP-ABE assay. Low-density cultures of embryonic rat hippocampal neurons have been widely used to study the localization, function, and trafficking of neuronal proteins, making them ideally suited for studying neuronal protein palmitoylation using the IP-ABE assay. The IP-ABE assay mainly requires standard IP and western blotting reagents, and is only limited by the availability of antibodies against the target substrate. This assay can easily be adapted for the purification and detection of transfected palmitoylated proteins in heterologous cell cultures, primary neuronal cultures derived from various brain tissues of both mouse and rat, and even primary brain tissue itself.

The IP-ABE assay specifically detects thiol-palmitoylation of cysteine residues along substrate proteins, and can be used to detect palmitoylation of immunoprecipitated neuronal proteins in a manner depicted in Figure 1. Treatment of the immunoprecipitated neuronal protein with HAM (Figure 1) cleaves the thioester linkage between palmitate and a cysteine's thiol group, allowing for specific incorporation of biotin-BMCC onto the newly available thiol group, which can be subsequently detected using western blotting. The omission of HAM (minus-HAM) cleavage prevents the incorporation of biotin-BMCC and functions as a negative control for specific palmitoyl-biotinylation of the plus-HAM sample, and should therefore always be run adjacent to the plus-HAM sample for western blotting by SDS-PAGE (Figure 2).
In an optimized experiment (Figure 2A), the palmitoylation signal of the neuronal protein &delta;-catenin2 can be readily detected by blotting for biotin-BMCC at a concentration of 1 &mu;M, using streptavidin conjugated with HRP, against parallel minus- and plus-HAM samples. The specific palmitoylation signal for &delta;-catenin appears at its predicted size of 160kD, only in the plus-HAM sample. The specificity of this signal was confirmed by reprobing for &delta;-catenin with the specific antibody that was initially used to immunoprecipitate it, resulting in a signal in both HAM treatments at the same predicted size. Optimization of the IP-ABE assay (Figure 2A) will result in a western blotting profile of a minus-HAM sample that is readily distinguishable from the plus-HAM sample, and exhibits minimal to no palmitoylation signal.
The concentration of biotin-BMCC used to label palmitoylated cysteines requires careful optimization, and if a super-saturating concentration of biotin-BMCC is used during the ABE chemistry steps (Figure 2B), excessive background and non-specific signals will be visible. A linear response curve for biotinylation of a palmitoylated neuronal protein, the &alpha;7 nicotinic acetylcholine receptor, using biotin-BMCC has previously been determined6, and shows near complete saturation at a concentration of 5-10 &mu;M. Although the optimal concentration of biotin-BMCC will deviate slightly depending on the neuronal target protein, treatment with biotin-BMCC at a concentration in excess of 5 &mu;M will likely super-saturate the target protein, and result in a western blot profile with visible background and non-specific signals. A concentration of 0.5 &mu;M for biotin-BMCC was previously used to detect palmitoylation of other neuronal proteins including SNAP-25, huntingtin, AMPA receptor subunits GluA1 and GluA2, and paralemmin8, and determination of the optimal concentration for a novel target protein can be accomplished by trial and error in linear fashion, starting within the range of 0.5-5 &mu;M that we describe here. The resulting streptavidin western blot profiles among the minus- and plus-HAM samples for &delta;-catenin palmitoylation appear similar when treated with 4 &mu;M biotin-BMCC (Figure 2B), with additional background signals at different sizes, which indicates that inappropriate biotinylation occurred.
Hydroxylamine is a powerful reducing agent that results in limited degradation of the target protein in addition to its efficient cleavage of thioester linkage. Consequently, optimization of ABE chemistry necessitates one to normalize the amount of immunoprecipitated protein used for minus- and plus-HAM samples (steps 3.2 - 3.3). Normalization requires the use of double the amount of immobilized target protein in the plus- vs. the minus-HAM sample, which actually results in indistinguishable western blotting profiles for the target protein, as shown for &delta;-catenin (Figure 2A, B). However, if the amount of immobilized target protein is not normalized between minus- and plus-HAM samples, the resultant western blot signal for the target protein in the plus-HAM sample can be lost, as shown for &delta;-catenin when equal amounts of protein were used in both HAM treatments (Figure 2C).
The specificity of ABE chemistry for labeling palmitoylated proteins is very sensitive and requires optimization. The common pitfalls encountered during the IP-ABE assay include the sub-optimal results shown in Figures 2B and 2C, and degradation of the target protein, independent of HAM treatment, which are typically caused by older reagents and buffers, and incorrect pH. In order to avoid these pitfalls and correct for sub-optimal ABE chemistry, the freshness and concentration of the reagents required for the buffers listed in Table 1 should always be examined, and the pH adjustments of all the buffers should always be checked prior to running the experiment.
<table border="1">
 <tbody>
 <tr>
 <td colspan="2">Buffer</td>
 <td>Working Concentration</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Lysis Buffer (LB)</td>
 <td>In distilled H2O: 1% IGEPAL CA-630 50mM Tris-HCl pH7.5 150mM NaCl 10% Glycerol</td>
 <td>N/A</td>
 <td>Prepare before experiment and store at 4 &deg;C</td>
 </tr>
 <tr>
 <td>NEM Solution</td>
 <td>In 100% EtOH: NEM lyophilized powder</td>
 <td>2 M</td>
 <td>Prepare fresh, immediately before use.</td>
 </tr>
 <tr>
 <td>Stringent Buffer</td>
 <td>In LB: 10 mM MEM 0.1% SDS</td>
 <td>N/A</td>
 <td>Prepare shortly before use, keep on ice.</td>
 </tr>
 <tr>
 <td>LB pH 7.2</td>
 <td>In LB: Adjust PH to exactly 7.2</td>
 <td>N/A</td>
 <td>Use a pH meter to adjust pH immediately before use.</td>
 </tr>
 <tr>
 <td>HAM Buffer</td>
 <td>In LB pH 7.2: Stock HAM solution</td>
 <td>1 M (final HAM concentration)</td>
 <td>Prepare immediately before use.</td>
 </tr>
 <tr>
 <td>LB pH 6.2</td>
 <td>In LB: Adjust pH to exactly 6.2</td>
 <td>N/A</td>
 <td>Same as for LB pH 7.2</td>
 </tr>
 <tr>
 <td>Stock Biotin-BMCC Solution</td>
 <td>In DMSO: 2.1 mg Biotin-BMCC solution</td>
 <td>8 mM</td>
 <td>Prepare immediately before use.</td>
 </tr>
 <tr>
 <td>Biotin-BMCC Buffer</td>
 <td>In LB pH 6.2: Add Biotin-BMCC solution</td>
 <td>1 - 5 &mu;M (final biotin-BMCC concentration)</td>
 <td>Prepare immediately before use.</td>
 </tr>
 <tr>
 <td>2x SDS Sample Buffer, no reducing agents</td>
 <td>In distilled H2O: 5% SDS 5% Glycerol 125mM Tris-HCL pH 6.8 0.01% Bromophenol Blue</td>
 <td>N/A</td>
 <td>Prepare before experiment, and store at -20 &deg;C until use. Supplement with 5 mM DTT before use.</td>
 </tr>
 </tbody>
</table>
Table 1. Buffers and reagents required for the IP-ABE assay. Many of the lysis buffers can be prepared before beginning the experiment, however the pH should be checked and adjusted immediately before use with a pH meter, to ensure success of the ABE chemistry. The majority of stock solutions should be prepared fresh for every experiment, immediately before use. Many of the buffers require reagents common to most laboratories equipped for biochemistry, and specialty reagents with vendor information are listed in the table of reagents and equipment.
<img src="/files/ftp_upload/50031/50031fig1.jpg" alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/50031/50031fig1highres.jpg" /> 
 Figure 1. Schematic of the Immunoprecipitation and Acyl-Biotin Exchange (IP-ABE) assay to purify and detect palmitoylation of neuronal proteins. Cultured hippocampal neurons are lysed, (1) a target protein is then purified using a target-specific antibody, and immobilized on sepharose beads coated with protein G or A. The purified target protein is then (2) treated with N-ethylmaliemide (NEM) to irreversibly bind and block free thiol (-SH) groups along unmodified cysteines (C). The target protein is then (3) subjected to treatment with hydroxylamine (HAM), resulting in specific cleavage of thioester bonds at palmitoylated cysteines and the unmasking of a free palmitoylated thiol group (-SH). Next, the target protein is (4) treated with a thiol-reactive biotin molecule, biotin-BMCC, resulting in specific biotinylation of the palmitoylated cysteine. Finally, (5) the biotinylated target protein is eluted and removed from the antibody and bead. The target protein with its palmitoylated cysteine(s) tagged with biotin is now suitable for SDS poly-acrylamide gel electrophoresis (SDS-PAGE), and western blotting with streptavidin to detect for palmitoylation of the purified neuronal protein. <a href="https://www.jove.com/files/ftp_upload/50031/50031fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/50031/50031fig2.jpg" alt="Figure 2" fo:content-width="6in" fo:src="/files/ftp_upload/50031/50031fig2highres.jpg"/> 
 Figure 2. Detection of palmitoylation of the neuronal protein &delta;-catenin using the IP-ABE assay. Primary rat hippocampal neurons were grown as previously described5, at a density of 130 cells/mm2 until maturity at 14DIV, were then lysed, and subjected to the IP-ABE assay to detect palmitoylation of a recently identified palmitoylated neuronal substrate, &delta;-catenin2. Purified immunoprecipitates (IPs) of &delta;-catenin (target protein) were isolated using 5 &mu;g of &delta;-catenin antibody per sample (BD Transduction Laboratories), and were then subjected to SDS-PAGE in parallel minus- and plus-hydroxylamine (HAM) samples. Palmitoylation was detected by western blotting (WB) with streptavidin-HRP (palmitoylation), and the membrane was stripped and subjected to a reprobe WB for &delta;-catenin. (A) An optimized representative IP-ABE result for palmitoylated &delta;-catenin (arrowhead) treated with 1 &mu;M biotin-BMCC, illustrating the specificity of ABE chemistry for detecting thiol-palmitoylated proteins from IP samples. (B) A sub-optimal representative IP-ABE result for &delta;-catenin, where an excess concentration of 4 &mu;M biotin-BMCC was used, resulting in non-specific signals and background in both minus- and plus-HAM samples (arrows). (C) Another sub-optimal representative IP-ABE WB for &delta;-catenin, where an excessive 4 &mu;M concentration of biotin-BMCC was used, and the amount of protein used for the plus-HAM IP sample was not normalized for HAM-mediated protein degradation, resulting in a lost signal in the reprobe WB for &delta;-catenin. <a href="https://www.jove.com/files/ftp_upload/50031/50031fig2large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE at JoVE.com

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

The reversible addition of palmitate to proteins is an important regulator of intracellular protein trafficking. This is of particular interest in neurons where many synaptic proteins are palmitoylated. We utilize a simple biochemical method to detect palmitoylated proteins in cultured neurons, which can be adapted for multiple cell types and tissues.

Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange (ABE)

Palmitoylation is a post-translational lipid  modification involving the attachment of a 16-carbon saturated fatty acid,  palmitate, to cysteine residues ...

detection-protein-palmitoylation-cultured-hippocampal-neurons

Research

JoVE Journal

Neuroscience

35.6K Aufrufe.  University of British Columbia. Die reversible Zugabe von Palmitat an Proteine ​​ist ein wichtiger Regulator des intrazellulären Protein-Trafficking. Dies ist von besonderem Interesse, wo viele Neuronen synaptische Proteine ​​palmitoyliert. Wir nutzen einen einfachen biochemischen Verfahren zur palmitoylierten Proteinen in kultivierten Neuronen, die für mehrere Zelltypen und Gewebe angepasst werden kann zu detektieren.

Serie: Detection of Protein Palmitoylation in Cultured Hippocampal Neurons by Immunoprecipitation and Acyl-Biotin Exchange ABE

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

Inducing Dendritic Growth in Cultured Sympathetic Neurons

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs 4-6. Altered patterns of dendritic growth and plasticity are associated with impaired neurobehavioral function in experimental models 7, and are thought to contribute to clinical symptoms observed in both neurodevelopmental disorders 8-10 and neurodegenerative diseases 11-13. Such observations underscore the functional importance of precisely regulating dendritic morphology, and suggest that identifying mechanisms that control dendritic growth will not only advance understanding of how neuronal connectivity is regulated during normal development, but may also provide insight on novel therapeutic strategies for diverse neurological diseases.
Mechanistic studies of dendritic growth would be greatly facilitated by the availability of a model system that allows neurons to be experimentally switched from a state in which they do not extend dendrites to one in which they elaborate a dendritic arbor comparable to that of their in vivo counterparts. Primary cultures of sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rodents provide such a model. When cultured in defined medium in the absence of serum and ganglionic glial cells, sympathetic neurons extend a single process which is axonal, and this unipolar state persists for weeks to months in culture 14,15. However, the addition of either bone morphogenetic protein-7 (BMP-7) 16,17 or Matrigel 18 to the culture medium triggers these neurons to extend multiple processes that meet the morphologic, biochemical and functional criteria for dendrites. Sympathetic neurons dissociated from the SCG of perinatal rodents and grown under defined conditions are a homogenous population of neurons 19 that respond uniformly to the dendrite-promoting activity of Matrigel, BMP-7 and other BMPs of the decapentaplegic (dpp) and 60A subfamilies 17,18,20,21. Importantly, Matrigel- and BMP-induced dendrite formation occurs in the absence of changes in cell survival or axonal growth 17,18.
Here, we describe how to set up dissociated cultures of sympathetic neurons derived from the SCG of perinatal rats so that they are responsive to the selective dendrite-promoting activity of Matrigel or BMPs.

We describe a protocol for using bone morphogenetic protein-7 (BMP-7) or Matrigel to selectively induce dendritic growth in primary sympathetic neurons dissociated from the superior cervical ganglia (SCG) of perinatal rats.

The shape of the dendritic arbor determines the total synaptic input a neuron can receive 1-3, and influences the types and distribution of these inputs ...

Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual neurons, researchers are able to analyze properties related to cellular trafficking, cellular structure and individual protein localization using a variety of biochemical techniques. Results from such experiments are critical for testing theories addressing the neural basis of memory and learning. However, unambiguous results from these forms of experiments are predicated on the ability to grow neuronal cultures with minimum contamination by other brain cell types. In this protocol, we use specific media designed for neuron growth and careful dissection of embryonic hippocampal tissue to optimize growth of healthy neurons while minimizing contaminating cell types (i.e. astrocytes). Embryonic mouse hippocampal tissue can be more difficult to isolate than similar rodent tissue due to the size of the sample for dissection. We show detailed dissection techniques of hippocampus from embryonic day 19 (E19) mouse pups. Once hippocampal tissue is isolated, gentle dissociation of neuronal cells is achieved with a dilute concentration of trypsin and mechanical disruption designed to separate cells from connective tissue while providing minimum damage to individual cells. A detailed description of how to prepare pipettes to be used in the disruption is included. Optimal plating densities are provided for immuno-fluorescence protocols to maximize successful cell culture. The protocol provides a fast (approximately 2 hr) and efficient technique for the culture of neuronal cells from mouse hippocampal tissue.

We provide a protocol for the culture of highly purified hippocampal neurons from prenatal mouse brains without the use of a feeder glial cell layer.

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Stripe Assay to Study the Attractive or Repulsive Activity of a Protein Substrate Using Dissociated Hippocampal Neurons

Growing axons develop a highly motile structure at their tip, termed the growth cone. The growth cone contacts extracellular environmental cues to navigate axonal growth. Netrin, slit, semaphorin, and ephrins are known guidance molecules that can attract or repel axons upon binding to receptors and co-receptors on the axon. The activated receptors initiate various signaling molecules in the growth cone that alter the structure and movement of the neuron. Here, we describe the detailed protocol for a stripe assay to assess the ability of a guidance molecule to attract or repel neurons. In this method, dissociated hippocampal neurons from E15.5 mice are cultured on laminin-coated dishes processed with alternating stripes of ectodomain of fibronectin and leucine-rich transmembrane protein-2 (FLRT2) and control immunoglobulin G (IgG) fragment crystallizable region (Fc) protein. Both axons and cell bodies were strongly repelled from the FLRT2-coated stripe regions after 24 h of culture. Immunostaining with tau1 showed that ~90% of the neurons were distributed on the Fc-coated stripes compared to the FLRT2-Fc-coated stripes (~10%). This result indicates that FLRT2 has a strong repulsive effect on these neurons. This powerful method is applicable not only for primary cultured neurons but also for a variety of other cells, such as neuroblasts.

Axon guidance molecules regulate neuronal migration and targeted growth-cone navigation. We present a powerful method, the stripe assay, to assess the ability of guidance molecules to attract or repulse neurons. In this protocol, we demonstrate the stripe assay by showing FLRT2's ability to repel cultured hippocampal neurons.