This work was supported by National Institutes of Health Grants NS-38849 and DK-076160, the Juvenile Diabetes Research Foundation Center for the Study of Complications in Diabetes, the Program for Neurology Research and Discovery and The A. Alfred Taubman Medical Research Institute at the University of Michigan. This work used the Morphology and Image Analysis Core of the Michigan Diabetes Research and Training Center, funded by National Institutes of Health Grant 5P60 DK-20572 from the National Institute of Diabetes and Digestive and Kidney Diseases. The authors thank Scott T. Clarke from Molecular Probes/Invitrogen for his valuable advice on the various Click-iT EdU labeling kits and the generous donation of reagents to support the initial development of the EdU amplification technique.

1. Preparation of Neurons
<ol>
 <li>Dorsal root ganglion (DRG) neurons are grown on sterile (autoclaved) 12 mm glass coverslips in a 24-well culture plate.</li>
 <li>The 10 mM stock of EdU (in DMSO, Click-iT EdU Microplate Assay Kit) is typically diluted 1:100 in culture medium to make a 10X EdU solution (100 &mu;M) and then diluted 1:10 into the culture wells (e.g. 30 &mu;L of the 10X EdU solution into a total of 300 &mu;L of culture medium). DRG neurons are incubated with a final concentration of 10 &mu;M EdU at 37&deg;C and 5% CO2 between 2-24 hours. The length of time depends on how treatments affect the rate of mtDNA synthesis.</li>
 <li>In a fume hood, DRG neurons are fixed in 2% paraformaldehyde for 10-15 min at room temperature, washed twice in 1X PBS 2-5 minutes each wash and stored in fresh 1X PBS for up to 1 month at 4&deg;C.</li>
 <li>Coverslips are transferred with fine-tipped forceps to a prepared humid chamber (Corning BioAssay dish with black papered bottom for contrast, wrapped in aluminum foil to protect light sensitive components and humidified with water saturated Kimwipes) and placed on a sheet of Parafilm M that provides a hydrophobic surface to confine the solutions to the coverslip. The protocol below is designed for 28 coverslips and solutions are prepared for 32 coverslips (about 14% extra volume). Solutions with expensive or limited components are made so that 75-80 &mu;L are used on each coverslip. Otherwise, 200-300 &mu;L are used for wash and block solutions to thoroughly wash out previous solutions.</li>
 <li>Re-apply 1X PBS to cover the surface of each coverslip (200-300 &mu;L). Prepare the Click-iT assay and tyramide signal amplification kits as described below and in the manufacturer's instructions.</li>
</ol>
Preparation of Click-iT EdU Microplate Assay Kit
Most components from the Click-iT EdU Microplate Assay Kit come pre-made and are stored at 4&deg;C [2x Click-iT Reaction Buffer (10x Component E), CuSO4 (100 mM, Component F), Click-iT EdU fixative (Component D) and Blocking Buffer (2x Component H)]. Click-iT EdU Buffer Additive (10x Component G) is stored at -20&deg;C to prevent it from turning yellow-brown over time. This component tolerates repeated freeze-thaw cycles.
The Oregon Green 488 azide (Component B) should be divided into small aliquots (10-20 &mu;L) to minimize freeze-thaw cycles and stored at -20&deg;C.
To prepare a stock solution of the anti-Oregon Green HRP conjugate (Component I), add 75 &mu;L of Milli Q dH2O to the vial. Mix by gentle pipetting or by inversion to avoid foaming and store at 4&deg;C. Do not vortex.
Preparation of TSA Kit #12, with HRP Goat Anti-Rabbit IgG and Alexa Fluor 488 Tyramide Kit
To prepare the tyramide stock solution, dissolve the solid material (Alexa Fluor 488 tyramide, Component A) in 150 &mu;L of DMSO (Component B). Invert the vial several times to dissolve any tyramide coating the sides of the vial. Store stock solution in small aliquots (10-20 &mu;L) at &le;-20&deg;C, desiccated and protected from light.
2. Click-iT 5-ethynyl-2-deoxyuridine (EdU) Labeling
NOTE: All solutions are removed with a bulb transfer pipette with a 200 &mu;L tip affixed to the end to gently remove liquids without losing cells. Avoid using a vacuum line, which usually remove solutions too vigorously. A bulb pipette is used to gently flood coverslips with 200-300 &mu;L of wash solutions to thoroughly wash out the previous solution.
<ol>
 <li>Cells are permeabilized with 0.1% Triton-X-100 in 1X PBS solution for 10 minutes at room temperature in a covered humid chamber. Use 1% Triton-X-100 stock solutions.
 <ul>
 <li>Make 3000 &mu;L of 0.1% Triton-X-100 by adding 300 &mu;L 1% Triton-X-100 stock to 2700&mu;L 1X PBS.</li>
 <li>Use 75-80 &mu;L per coverslip.</li>
 </ul>
 </li>
 <li>Remove the Triton X-100 solution and rinse twice with 1X PBS.</li>
 <li>Endogenous peroxidase activity is quenched by a 1% H2O2 in 1X PBS solution for 30 minutes at room temperature. Dilute 30% H2O2 solution with 1X PBS. This solution should be made fresh but can be made during the 10 minute permeabilization step above (step 3.1).
 <ul>
 <li>Make 3000 &mu;L of a 1% H2O2 by adding 100 &mu;L 30% H2O2 to 2900 &mu;L 1X PBS.</li>
 <li>Use 75-80 &mu;L per coverslip.</li>
 <li>Remove the peroxidase solution and rinse twice with 1X PBS.</li>
 </ul>
 </li>
 <li>Click-iT EdU Reaction
 <ul>
 <li>For 32 coverslips (32 x 80 &mu;L = 2560 &mu;L) a total of 2560 &mu;L is needed. The 2x reaction is made in 1280 &mu;L, which is half of the total volume of the reaction.</li>
 <li>Freshly prepare the Click-iT Reaction Cocktail prior to starting the 5 minute post fix (step 3.5 below).</li>
 <li>Mix the cocktail by pipetting up and down. Do not vortex when making this reaction. 
 <table border="1" cellpadding="0" cellspacing="0">
 <tbody>
 <tr>
 <td>2 x Reaction Cocktail 
 Components are from Click-iT EdU Microplate Assay Kit</td>
 <td>Half Volume: 1280 &mu;L</td>
 </tr>
 <tr>
 <td>Milli Q dH2O</td>
 <td> 1132.8 &mu;L</td>
 </tr>
 <tr>
 <td>2x Click-iT Reaction Buffer (10x Component E)</td>
 <td> 100.3 &mu;L</td>
 </tr>
 <tr>
 <td>Click-iT EdU Buffer Additive (10x Component G)</td>
 <td> 25.6 &mu;L</td>
 </tr>
 <tr>
 <td>CuSO4 (100 mM, Component F)</td>
 <td> 25.6 &mu;L</td>
 </tr>
 <tr>
 <td>Oregon Green Azide (Component A)</td>
 <td> 6.4 &mu;L</td>
 </tr>
 <tr>
 <td>Total</td>
 <td>1290.7 &mu;L</td>
 </tr>
 </tbody>
 </table>
 
 NOTE: The volumes used above are proportional to the volumes listed in the Click-iT EdU Microplate Assay kit directions. The final volume of Reaction Cocktail is slightly more than 1280 &mu;L.</li>
 <li>The 2X Click-iT reaction is diluted with equal volume of Click-iT EdU fixative (Component D) right before use. Mixing these together makes a uniform reaction solution for all coverslips.</li>
 <li>Add 1290.7 &mu;L Click-iT EdU fixative (Component D) to the 1290.7 &mu;L reaction cocktail. Use 75-80 &mu;L per coverslip.</li>
 </ul>
 </li>
 <li>Post-fix the neurons with Click-iT EdU fixative (Component D) for 5 minutes at room temperature.</li>
 <li>Remove fix and add reaction cocktail from above (step 3.4). Cover to protect from light and incubate for 25 minutes at room temperature. Gently remove the Reaction Cocktail. Wash twice with diluted 1X Blocking Buffer (2X Component H).
 <ul>
 <li>Make 5200 &mu;L of 1X Blocking Buffer by diluting 2600 &mu;L Blocking Buffer (2x Component H) with an equal volume of MilliQ d H2O (2600 &mu;L). Use 75-80 &mu;L per coverslip.</li>
 </ul>
 </li>
 <li>Under an epi-fluorescent microscope, check for Oregon Green staining in nuclei of mitotically active control cells.</li>
</ol>
3. Tyramide Signal Amplification (TSA) of EdU Signal
<ol>
 <li>Add 1% TSA block solution to each coverslip and incubate for 30 minutes at room temperature.
 <ul>
 <li>Make 6000 &mu;L of 1% TSA block solution by weighing 0.06 g of TSA Blocking Reagent (TSA kit #12, Component D) and adding it to 6000 &mu;L 1X PBS. Vortex to mix. Addition of 5% goat serum in the 1% TSA block solution will often help to reduce non-specific binding.</li>
 </ul>
 </li>
 <li>Briefly spin the anti-Oregon Green horseradish peroxidase conjugated antibody stock (Component I of the Click-iT EdU Microplate Assay), prepared 2-24 hours in advance of TSA. Dilute the primary antibody 1:300 in 1% TSA blocking solution and invert or pipette up and down gently to mix. Do not vortex to avoid disrupting the HRP-conjugated antibody. Remove 1% TSA block solution, add 75 &mu;L to each coverslip and incubate overnight at 4&deg;C.
 <ul>
 <li>The antibody can be used more concentrated, such as 1:150, but it is supplied in limited amounts, so use it sparingly. Again, addition of 5% goat serum in the 1% TSA block solution will often help to reduce non-specific binding of the anti-Oregon Green horseradish peroxidase conjugated antibody.</li>
 <li>Make 2560 &mu;L of a 1:300 antibody solution by adding 8.53 &mu;L of the stock Rabbit anti-Oregon Green-HRP (Click-iT EdU Microplate Assay) to 2560 &mu;L 1% TSA Blocking. Mix by gentle pipetting or inversion.</li>
 </ul>
 </li>
 <li>The next day, remove the antibody solution and rinse three times with 1X PBS. Incubate the coverslips in the final wash for an additional 30-60 minutes at room temperature to ensure that unbound primary antibody is removed.</li>
 <li>Prepare Tyramide reaction for 2560 &mu;L (32 coverslips x 80 &mu;L).
 <ul>
 <li>Make 400 &mu;L of a 0.15% H2O2 solution (100X) by adding 2 &mu;L of the 30 % H2O2 (TSA kit #12, Component F) to 398 &mu;L Amplification Buffer (TSA kit #12, Component E). This should be made right before needed.</li>
 <li>For a final volume of 2560 &mu;L combine:
 <ul>
 <li>25.6 &mu;L Tyramide-488 (TSA kit Component A)</li>
 <li>2508.8 &mu;L Amplification Buffer (TSA kit Component E)</li>
 <li>25.6 &mu;L 0.15% H2O2 (from above for a final 0.0015% H2O2)</li>
 <li>Remove 1X PBS and incubate with the Tyramide reaction for 15 minutes at room temperature.</li>
 </ul>
 </li>
 </ul>
 </li>
 <li>Remove Tyramide reaction and rinse three times with 1X PBS, incubating in the final wash for 30-60 minutes at room temperature as before.</li>
 <li>Check for mtDNA labeling under an epi-fluorescent microscope.
 <ul>
 <li>Mount coverslips on glass microscope slides with an antifade mounting medium, such as ProLong Gold with DAPI. Alternatively, the EdU labeling and amplification can be followed by standard fluorescent immunocytochemistry to label other cellular markers.</li>
 </ul>
 </li>
</ol>
4. Representative Results: Visualization of Mitochondrial DNA Replication as a Marker for Mitochondrial Biogenesis
We are interested in how sensory neurons (Figure 1) regulate the number of mitochondria. This protocol will label newly synthesized mtDNA with a fluorescent marker as a way of measuring new mitochondria. The incorporation of the synthetic nucleotide EdU followed by the click chemistry of the Oregon Green-azide and subsequent amplification with the Alexa-Fluor 488 tyramide results in the labeling of mtDNA with a green fluorescent signal (Figure 2). If done correctly, the amplified green signal is sufficiently higher than background fluorescence (such as cellular autofluorescence or a side-effect of the HRP-TSA reaction).
This technique is designed to label newly replicated mtDNA in order to visualize and quantify mtDNA biogenesis within subcellular compartments of neurons (Figure 3). The EdU labeling allows for subsequent fluorescent immunocytochemistry to label other cellular markers such as neurofilament (Figure 3D).
The TSA reaction can also be done with other fluorescent Alexa Fluor tyramides such as Alexa Fluor 594-tyramide. This results in a green fluorescent nuclear signal from the Oregon Green-azide click reaction in the nucleus but no green signal in mtDNA, which is undetectable prior to TSA. The amplification with Alexa Fluor 594-tyramide intensifies the nuclear label and reveals the incorporated EdU in mtDNA with a red fluorescent signal (Figure 4). A similar amplification procedure is used to visualize BrdU incorporation into newly synthesized mtDNA (Figure 5), however, this method requires an additional step to recover the BrdU epitope by either a harsh acid (HCl) or enzyme digest, which is not necessary for EdU labeling.
<img src="/files/ftp_upload/2147/2147fig1.jpg" alt="Figure 1" /> 
 Figure 1. Representative differential interference contrast (DIC) images of embryonic (left) and adult (right) dorsal root ganglion (DRG) neurons that are typically used for analysis. Bars = 10 &mu;m.
<img src="/files/ftp_upload/2147/2147fig2.jpg" alt="Figure 2" /> 
 Figure 2. Schematic diagram representing the procedure for labeling EdU in mtDNA with a green fluorescent signal. The schematic represents the three-step protocol for labeling EdU in mtDNA with a green fluorescent signal. Mitotically active F11 neuroblastoma cells serve as a positive control and illustrate the labeling pattern of EdU that was incorporated in nuclear and mitochondrial DNA. The first step (P1) is to incorporate a thymidine analog into newly synthesized mtDNA by incubating cells in the presence of EdU. The second step (P2) is based on click chemistry to label incorporated EdU with an Oregon Green-azide. Green signal is visible in nuclei of cells that replicated their nuclear DNA. The final step (P3) is to amplify the Oregon Green-azide signal by incubating with a HRP-conjugated rabbit antibody against Oregon Green followed by incubation with Alexa Fluor 488 labeled tyramide in the presence of hydrogen peroxide (H2O2) to visualize green signal in mtDNA.
<img src="/files/ftp_upload/2147/2147fig3.jpg" alt="Figure 3" /> 
 Figure 3. EdU labeling of mtDNA in dorsal root ganglion (DRG) neurons. Neurons are incubated with EdU and the signal is subsequently amplified to reveal mtDNA that incorporated EdU. Representative fluorescence images overlaid on transmitted light showing green punctate signals of amplified EdU (EdU-OG-TSA, green) incorporated into newly synthesized mtDNA of both embryonic (A) and adult (B-D) DRG neurons. The EdU labeling procedure allows for subsequent immunofluorescence staining of neuronal markers such as neurofilament (D, &alpha;NF, in red). Scale bars = 10 &mu;m.
<img src="/files/ftp_upload/2147/2147fig4.jpg" alt="Figure 4" /> 
 Figure 4. Schematic diagram representing the procedure for labeling EdU in mtDNA with a red fluorescent signal. The schematic represents the three-step protocol for labeling EdU in mtDNA with a red fluorescent signal. Mitotically active F11 neuroblastoma cells serve as a positive control and illustrate the labeling pattern of EdU that was incorporated in nuclear and mitochondrial DNA. The first step (P1) is to incorporate a thymidine analog into newly synthesized mtDNA by incubating cells in the presence of EdU. The second step (P2) is based on click chemistry to label incorporated EdU with an Oregon Green-azide. Green signal is visible in nuclei of cells that replicated their nuclear DNA. The final step (P3) is to amplify the Oregon Green-azide signal by incubating with a HRP-conjugated rabbit antibody against Oregon Green followed by incubation with Alexa Fluor 594 labeled tyramide in the presence of hydrogen peroxide (H2O2) to visualize red signal in mtDNA.
<img src="/files/ftp_upload/2147/2147fig5.jpg" alt="Figure 5" /> 
 Figure 5. BrdU labeling and tyramide signal amplification with a green or red fluorescent signal in mtDNA. The schematic diagram represents the procedure for labeling BrdU in newly synthesized mtDNA with either a green or red fluorescent signal. An initial step is required to recover the BrdU epitope by either an acid (HCl) or enzyme digest, which is not necessary for EdU labeling. The next step is incubating with a mouse primary anti-BrdU antibody. This is followed by incubating with a horseradish peroxidase (HRP)-conjugated goat anti-mouse antibody. Finally, the signal is amplified with a green or red fluorescently labeled tyramide in the presence of hydrogen peroxide (H2O2) to visualize signal in mtDNA.

<ol>
	<li>Chan, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+fusion+and+fission+in+mammals.">Mitochondrial fusion and fission in mammals.</a> Annu. Rev. Cell Dev. Biol. 22, 79-99 (2006).</li><li>Dimmer, K. S., Scorrano, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=(De)constructing+mitochondria:+what+for.">(De)constructing mitochondria: what for.</a> Physiology (Bethesda). 21, 233-241 (2006).</li><li>Suen, D. F., Norris, K. L., Youle, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+dynamics+and+apoptosis.">Mitochondrial dynamics and apoptosis.</a> Genes Dev. 22, 1577-1590 (2008).</li><li>Clay Montier, L. L., Deng, J. J., Bai, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Number+matters:+control+of+mammalian+mitochondrial+DNA+copy+number.">Number matters: control of mammalian mitochondrial DNA copy number.</a> J. Genet. Genomics. 36 (3), 125-131 (2009).</li><li>Salic, A., Mitchison, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+chemical+method+for+fast+and+sensitive+detection+of+DNA+synthesis+in+vivo.">A chemical method for fast and sensitive detection of DNA synthesis in vivo.</a> Proc Natl Acad Sci U.S.A. 105, 2415-2420 (2008).</li><li>Buck, S. B., Bradford, J., Gee, K. R., Agnew, B. J., Clarke, S. T., Salic, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+S-phase+cell+cycle+progression+using+5-ethynyl-2'-deoxyuridine+incorporation+with+click+chemistry,+an+alternative+to+using+5-bromo-2'-deoxyuridine+antibodies.">Detection of S-phase cell cycle progression using 5-ethynyl-2'-deoxyuridine incorporation with click chemistry, an alternative to using 5-bromo-2'-deoxyuridine antibodies.</a> Biotechniques. 44, 927-929 (2008).</li><li>Yu, Y., Arora, A., Min, W., Roifman, C. M., Grunebaum, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EdU+incorporation+is+an+alternative+non-radioactive+assay+to+[(3)H]thymidine+uptake+for+in+vitro+measurement+of+mice+T-cell+proliferations.">EdU incorporation is an alternative non-radioactive assay to [(3)H]thymidine uptake for in vitro measurement of mice T-cell proliferations.</a> J Immunol Methods. 350 (1-2), 29-35 (2009).</li><li>Heusden, J. V. a. n., Jong, P. d. e., Ramaekers, F., Bruwiere, H., Borgers, M., Smets, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescein-labeled+tyramide+strongly+enhances+the+detection+of+low+bromodeoxyuridine+incorporation+levels.">Fluorescein-labeled tyramide strongly enhances the detection of low bromodeoxyuridine incorporation levels.</a> J. Histochem. Cytochem. 45 (2), 315-319 (1997).</li><li>Cappella, P., Gasparri, F., Pulici, M., Moll, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+based+on+click+chemistry,+which+overcomes+limitations+of+cell+cycle+analysis+by+classical+determination+of+BrdU+incorporation,+allowing+multiplex+antibody+staining.">A novel method based on click chemistry, which overcomes limitations of cell cycle analysis by classical determination of BrdU incorporation, allowing multiplex antibody staining.</a> Cytometry A. 73, 626-636 (2008).</li><li>Kaiser, C. L., Kamien, A. J., Shah, P. A., Chapman, B. J., Cotanche, 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=5-Ethynyl-2'-deoxyuridine+labeling+detects+proliferating+cells+in+the+regenerating+avian+cochlea.">5-Ethynyl-2'-deoxyuridine labeling detects proliferating cells in the regenerating avian cochlea.</a> Laryngoscope. 119, 1770-1775 (2009).</li><li>Lentz, S. I., Edwards, J. L., Backus, C., McLean, L. L., Haines, K. M., Feldman, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+DNA+(mtDNA)+Biogenesis:+Visualization+and+Duel+Incorporation+of+BrdU+and+EdU+Into+Newly+Synthesized+mtDNA+In.">Mitochondrial DNA (mtDNA) Biogenesis: Visualization and Duel Incorporation of BrdU and EdU Into Newly Synthesized mtDNA In.</a> Vitro. J Histochem Cytochem. 58 (2), 207-218 (2010).</li><li>Edwards, J. L., Quattrini, A., Lentz, S. I., Figueroa-Romero, C., Cerri, F., Backus, C., Hong, Y., Feldman, E. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetes+regulates+mitochondrial+biogenesis+and+fission+in+mouse+neurons.">Diabetes regulates mitochondrial biogenesis and fission in mouse neurons.</a> Diabetologia. 53 (1), 160-169 (2010).</li><li>Davis, A. F., Clayton, 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=In+situ+localization+of+mitochondrial+DNA+replication+in+intact+mammalian+cells.">In situ localization of mitochondrial DNA replication in intact mammalian cells.</a> J. Cell Biol. 135, 883-893 (1996).</li><li>Magnusson, J., Orth, M., Lestienne, P., Taanman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Replication+of+mitochondrial+DNA+occurs+throughout+the+mitochondria+of+cultured+human+cells.">Replication of mitochondrial DNA occurs throughout the mitochondria of cultured human cells.</a> Exp. Cell. Res. 289, 133-142 (2003).</li><li>Amiri, M., Hollenbeck, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+biogenesis+in+the+axons+of+vertebrate+peripheral+neurons.">Mitochondrial biogenesis in the axons of vertebrate peripheral neurons.</a> Dev. Neurobiol. 68, 1348-1361 (2008).</li></ol>

No conflicts of interest declared.

We developed a sensitive assay to label newly synthesized mtDNA in individual cells using a tyramide signal amplification of EdU. One of the biggest issues during the optimization of this protocol was the inconsistent results between coverslips. Changes were made to the Invitrogen kit protocols to avoid mixing small volumes on individual coverslips and making master solutions to use for all coverslips. The 75-80 &mu;L used for each 12 mm circular glass coverslip is an ideal volume to completely cover the surface area while providing enough solution to give consistent results between samples. Incubation times and reagent concentrations were optimized but improvements might be seen with adjustments to them.
The protocol is designed to run each process once. However, some of the steps have been repeated with new solutions to recover samples that failed after the first attempt. In particular, the EdU click reaction steps (3.4-3.6) have been repeated without a significant increase in background fluorescence. The anti-Oregon Green-HRP antibody incubation (4.1-4.3) and the tyramide amplification (4.4-4.5) steps have also been repeated, but more often results in a poor signal to noise ratio because of the increased background fluorescence.
The most sensitive reagents are the Click-iT EdU Buffer Additive (Component G) and the rabbit anti-Oregon Green-HRP antibody (Component I) from the Click-iT EdU Microplate Assay kit. The Click-iT EdU Buffer Additive gradually turns yellow over time when stored at 4&deg;C and between 6-12 months darkens considerably. Storage of the Click-iT EdU Buffer Additive at -20&deg;C will alleviate this issue. The labeling and tyramide amplification steps work best when the rabbit anti-Oregon Green-HRP antibody is used within 4-6 months of reconstituting it in MilliQ dH2O.
The mild labeling of EdU in mtDNA allows for direct comparisons with additional cellular markers9-11 and enhances the utility of this technique over other thymidine analogs, such as 5-bromo-2-deoxyuridine (BrdU), which requires a harsh treatment to recover its epitope within DNA. Our laboratory11-12 and others13-15 have successfully used BrdU to label mtDNA. The BrdU labeling technique is similar to the one used for EdU but has some important differences (Figure 5). After the permeabilization step listed above (3.1-3.2), the BrdU epitope is recovered with a denaturation step (2 N HCl for 30 min at 37&deg;C followed by three washes in 1X PBS). The BrdU is then labeled with a primary antibody against BrdU (Vector Laboratories diluted 1:50 in 1% blocking solution from the TSA kit and incubated overnight at 4&deg;C). A secondary antibody step is needed to amplify the BrdU signal (goat anti-mouse IgG conjugated to HRP from the TSA kits #2 and #5, diluted 1:100 in 1% blocking solution and incubated for 45 min at room temperature) prior to the TSA steps (4.3-4.5 above). Having two thymidine analogs, EdU and BrdU, to label mtDNA is advantageous for performing dual label experiments where mtDNA can be sequentially labeled in pulse-labeling paradigms11.
Our laboratory is using the EdU and BrdU labeling of mtDNA to examine the regulation of mitochondrial biogenesis in the context of diabetic neuropathy, a common complication of diabetes. We have successfully used the signal amplification to measure changes in mtDNA biogenesis in individual neurons11-12. This technique will be useful in other experiments designed to explore the mechanisms of mtDNA replication and turnover or for identifying drugs that inhibit mtDNA synthesis. In addition, the basic principles of amplifying EdU and BrdU signals may be applied to other studies that measure DNA replication or repair.

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		<div>
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		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>12 mm coverslips</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>12-545-82</td>
<td>&nbsp;</td>
<tr>
<td>245 mm x 245 mm BioAssay Dish</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>431111</td>
<td>&nbsp;</td>
<tr>
<td>Parafilm M</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>PM-996</td>
<td>&nbsp;</td>
<tr>
<td>paraformaldehye</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>P6148</td>
<td>&nbsp;</td>
<tr>
<td>Triton X-100</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>T-8532</td>
<td>&nbsp;</td>
<tr>
<td>Phosphate buffered saline 10X solution</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>BP399</td>
<td>&nbsp;</td>
<tr>
<td>Transfer Pipet</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>13-711-7M</td>
<td>&nbsp;</td>
<tr>
<td>Hydrogen peroxide, 30% in water</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>BP2633</td>
<td>&nbsp;</td>
<tr>
<td>Click-iT EdU Microplate Assay Kit</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>C10214</td>
<td>&nbsp;</td>
<tr>
<td>TSA Kit #12, with HRP&#x2014;goat anti-rabbit IgG and Alexa Fluor 488 tyramide</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>T20922</td>
<td>&nbsp;</td>
<tr>
<td>TSA Kit #15, with HRP&#x2013;goat anti-rabbit IgG and Alexa Fluor 594 tyramide</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>T20925</td>
<td>&nbsp;</td>
<tr>
<td>ProLong Gold antifade reagent with DAPI</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P-36931</td>
<td>&nbsp;</td>
<tr>
<td>Polyclonal rabbit Neurofilament antibody</td>
<td>&nbsp;</td>
<td>Chemicon</td>
<td>AB1981</td>
<td>&nbsp;</td>
<tr>
<td>Mouse monoclonal anti-BrdU antibody</td>
<td>&nbsp;</td>
<td>Vector Laboratories</td>
<td>VP-B209</td>
<td>&nbsp;</td>
<tr>
<td>TSA Kit #2, with HRP&#x2014;goat anti-mouse IgG and Alexa Fluor 488 tyramide</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>T20912</td>
<td>&nbsp;</td>
<tr>
<td>TSA Kit #5, with HRP&#x2014;goat anti-mouse IgG and Alexa Fluor 594 tyramide</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>T20915</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

visualization of mitochondrial dna replication in individual cells by edu signal amplification

Mitochondria are key regulators of cellular energy and mitochondrial biogenesis is an essential component of regulating mitochondria numbers in healthy cells1-3. One approach for monitoring mitochondrial biogenesis is to measure the rate of mitochondrial DNA (mtDNA) replication4. We developed a sensitive technique to label newly synthesized mtDNA in individual cells in order to study mtDNA biogenesis. The technique combines the incorporation of 5-ethynyl-2'-deoxyuridine (EdU)5-7 with a tyramide signal amplification (TSA)8 protocol to visualize mtDNA replication within subcellular compartments of neurons. EdU is superior to other thymidine analogs, such as 5-bromo-2-deoxyuridine (BrdU), because the initial click reaction to label EdU5-7 does not require the harsh acid treatments or enzyme digests that are required for exposing the BrdU epitope. The milder labeling of EdU allows for direct comparison of its incorporation with other cellular markers9-10. The ability to visualize and quantify mtDNA biogenesis provides an essential tool for investigating the mechanisms used to regulate mitochondrial biogenesis and would provide insight into the pathogenesis associated with drug toxicity, aging, cancer and neurodegenerative diseases. Our technique is applicable to sensory neurons as well as other cell types. The use of this technique to measure mtDNA biogenesis has significant implications in furthering the understanding of both normal cellular physiology as well as impaired disease states.

Watch this Scientific Journal Video about Visualization of Mitochondrial DNA Replication in Individual Cells by EdU Signal Amplification at JoVE.com

Visualization of Mitochondrial DNA Replication in Individual Cells by EdU Signal Amplification

We developed a sensitive technique to label newly synthesized mitochondrial DNA (mtDNA) in individual cells in order to study mtDNA biogenesis. The technique combines the incorporation of EdU together with a tyramide signal amplification (TSA) protocol to visualize mtDNA replication within subcellular compartments of neurons.

Mitochondria are key regulators of cellular energy and mitochondrial biogenesis is an essential component of regulating mitochondria numbers in healthy ...

visualization-mitochondrial-dna-replication-individual-cells-edu

Research

JoVE Journal

Neuroscience

22.9K 조회수.  University of Michigan. 우리는 mtDNA의 biogenesis를 공부하기 위해서는 각각의 세포에서 새로 합성된 DNA mitochondrial (mtDNA)를 라벨에 민감한 기술을 개발했습니다. 기술은 뉴런의 subcellular 구획 내에 mtDNA 복제를 시각화하는 tyramide 신호 증폭 (TSA) 프로토콜과 함께 에듀의 결합이 결합되어 있습니다.

비디오: Visualization of Mitochondrial DNA Replication in Individual Cells by EdU Signal Amplification

Automated Interactive Video Playback for Studies of Animal Communication

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, parameter-based computer animation offers the opportunity to independently manipulate any number of behavioral, morphological, or spectral characteristics in the context of realistic, moving images of animals on screen. A major limitation of conventional playback, however, is that the visual stimulus lacks the ability to interact with the live animal. Borrowing from video-game technology, we have created an automated, interactive system for video playback that controls animations in response to real-time signals from a video tracking system. We demonstrated this method by conducting mate-choice trials on female swordtail fish, Xiphophorus birchmanni. Females were given a simultaneous choice between a courting male conspecific and a courting male heterospecific (X. malinche) on opposite sides of an aquarium. The virtual male stimulus was programmed to track the horizontal position of the female, as courting males do in the wild. Mate-choice trials on wild-caught X. birchmanni females were used to validate the prototype's ability to effectively generate a realistic visual stimulus.

Video playback is a widely used technique in animal behavior. We created and evaluated a program that applies rules-based, interactive playback of 3-D computer animations in response to real-time, automated data on subject behavior.

Video playback is a widely-used technique for the controlled manipulation and presentation of visual signals in animal communication. In particular, ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, direction5-7, ocular dominance8,9 and binocular disparity9. Orientation selectivity is the most widely studied feature and a continuous map with a quasi-periodic layout for preferred orientation is present across the entire primary visual cortex10,11. Integrating the synaptic, cellular and network contributions that lead to stimulus selective responses in these functional maps requires the hybridization of imaging techniques that span sub-micron to millimeter spatial scales. With conventional intrinsic signal optical imaging, the overall layout of functional maps across the entire surface of the visual cortex can be determined12. The development of in vivo two-photon microscopy using calcium sensitive dyes enables one to determine the synaptic input arriving at individual dendritic spines13 or record activity simultaneously from hundreds of individual neuronal cell bodies6,14. Consequently, combining intrinsic signal imaging with the sub-micron spatial resolution of two-photon microscopy offers the possibility of determining exactly which dendritic segments and cells contribute to the micro-domain of any functional map in the neocortex. Here we demonstrate a high-yield method for rapidly obtaining a cortical orientation map and targeting a specific micro-domain in this functional map for labeling neurons with fluorescent dyes in a non-rodent mammal. With the same microscope used for two-photon imaging, we first generate an orientation map using intrinsic signal optical imaging. Then we show how to target a micro-domain of interest using a micropipette loaded with dye to either label a population of neuronal cell bodies or label a single neuron such that dendrites, spines and axons are visible in vivo. Our refinements over previous methods facilitate an examination of neuronal structure-function relationships with sub-cellular resolution in the framework of neocortical functional architectures.

A method is described for labeling neurons with fluorescent dyes in predetermined functional micro-domains of the neocortex. First, intrinsic signal optical imaging is used to obtain a functional map. Then two-photon microscopy is used to label and image neurons within a micro-domain of the map.

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Evaluation of Bioenergetic Function in Cerebral Vascular Endothelial Cells

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types that comprise the BBB; these cells have a very high-energy demand, which requires optimal mitochondrial function. In the case of disease or injury, the mitochondrial function in these cells can be altered, resulting in disease or&#160;the opening of the BBB. In this manuscript, we introduce a method to measure mitochondrial function in CVE cells by using whole, intact cells and a bioanalyzer. A mito-stress assay is used to challenge the cells that have been perturbed, either physically or chemically, and evaluate their bioenergetic function. Additionally, this method also provides a useful way to screen new therapeutics that have direct effects on mitochondrial function. We have optimized the cell density necessary to yield oxygen consumption rates that allow for the calculation of a variety of mitochondrial parameters, including ATP production, maximal respiration, and spare capacity. We also show the sensitivity of the assay by demonstrating that the introduction of the&#160;microRNA, miR-34a, leads to a pronounced and detectable decrease in mitochondrial activity. While the data shown in this paper is optimized for the bEnd.3 cell line, we have also optimized the protocol for primary CVE cells, further suggesting the utility in preclinical and clinical models.

Endothelial cell mitochondria are critical to maintain blood-brain-barrier integrity. We introduce a protocol to measure bioenergetic function in cerebral vascular endothelial cells.

The integrity of the blood-brain-barrier (BBB) is critical to prevent brain injury. Cerebral vascular endothelial (CVE) cells are one of the cell types ...

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

The production of ATP by oxidative phosphorylation is the primary function of mitochondria. Mitochondria in higher eukaryotes also participate in cytosolic Ca2+ buffering, and the ATP production in mitochondrial can be mediated by intramitochondrial free Ca2+ concentration. Ca2+ retention capacity can be regarded as the capability of mitochondria to retain calcium in the mitochondrial matrix. Accumulated intracellular Ca2+ leads to the permeability of the inner mitochondrial membrane, termed the opening of mitochondrial permeability transition pore (mPTP), which leads to the leakage of molecules with a molecular weight less than 1.5 kDa. Ca2+-triggered mitochondria swelling is used to indicate the mPTP opening. Here, we describe two assays to examine the Ca2+ retention capacity and Ca2+-triggered mitochondrial swelling in isolated mitochondria. After certain amounts of Ca2+ are added, all steps can be completed in one day and recorded by a microplate reader. Thus, these two simple and effective assays can be adopted to assess the Ca2+-related mitochondrial functions.

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

This protocol aims to describe a method to examine the Ca2+ retention capacity and Ca2+- triggered mitochondrial swelling of isolated mitochondria of SH-SY5Y cells step-by-step.

The production of ATP by oxidative phosphorylation is the primary function of mitochondria. Mitochondria in higher eukaryotes also participate in ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

Visualization of Cortical Modules in Flattened Mammalian Cortices

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and can be delineated by certain histochemical and immunohistochemical methods. In this study, we highlight a method to isolate the cortex from mammalian brains and flatten them to obtain sections parallel to the cortical sheet. We further highlight selected histochemical and immunohistochemical methods to process these flattened tangential sections to visualize cortical modules. In the somatosensory cortex of various mammals, we perform cytochrome oxidase histochemistry to reveal body maps or cortical modules representing different parts of the body of the animal. In the medial entorhinal cortex, an area where grid cells are generated, we utilize immunohistochemical methods to highlight modules of genetically determined neurons which are arranged in a grid-pattern in the cortical sheet across several species. Overall, we provide a framework to isolate and prepare layer-wise flattened cortical sections, and visualize cortical modules using histochemical and immunohistochemical methods in a wide variety of mammalian brains.

This article describes a detailed methodology to obtain flattened tangential sections from mammalian cortices and visualize cortical modules using histochemical and immunohistochemical methods.

The cortex of mammalian brains is parcellated into distinct substructures or modules. Cortical modules typically lie parallel to the cortical sheet, and ...