We thank Michael Candlish for critical comments on the manuscript. This project was supported by the Deutsche Forschungsgemeinschaft grants BO1743/6 and SFB/TRR 152 P11 and Z02 to Ulrich Boehm.

NOTE: Ethics Statement: Procedures involving animal subjects were approved by the animal welfare committee of the University of Hamburg and the University of Saarland.
1. Preparation and Fixation of Embryonic Tissue
<ol>
	<li>Arrange all the equipment needed to dissect out the embryos and prepare solutions for subsequent fixation of the tissue before sacrificing the animals. 
	NOTE: Always prepare a fresh 4% paraformaldehyde (PFA) solution (4% PFA in 0.1 M phosphate buffered saline (PBS), adjust pH to 7.4).</li>
	<li>Euthanize timed pregnant female mouse with an ethically approved procedure.</li>
	<li>Transfer the mouse onto a platform. Wet the ventral side of the abdomen with 70% ethanol and make a vertical incision using sharp scissors to expose the abdominal cavity.</li>
	<li>Identify the embryonic chain and pull it out carefully using blunt forceps and place it into ice-cold PBS.</li>
	<li>Remove the individual embryos from the embryonic chain using fine scissors and forceps. Remove extra tissue around the embryos and wash them in ice-cold PBS.</li>
	<li>Take a small tail biopsy (before transferring the embryo into PFA) and incubate it in tail lysis buffer for the gender identification and tracer allele genotyping PCR.</li>
	<li>Start from the embryo at the most lateral side and transfer each embryo separately into a tube containing ice-cold 4% PFA on ice on a shaker.</li>
	<li>Soak the embryos at the age of E13.5 or younger for 1.5 hr on ice with constant shaking. Soak the embryos at age E14.5 and E15.5 for 2.5 hr on ice with constant shaking.</li>
	<li>For embryos age E16.5 or older, decapitate and remove the outer skin before soaking the head in PFA solution. Heads of E16.5 embryos can be soaked for 4.5 hr on ice with constant shaking.</li>
	<li>After appropriate fixation, wash the embryos three times with ice-cold PBS. Transfer the embryos into 30% sucrose in PBS solution at 4 &#176;C until they sink to the bottom.</li>
</ol>
2. Freezing
<ol>
	<li>Arrange all the materials required before starting the freezing process: cryo-mold, cryo-gloves, featherweight entomology forceps, marker pen and optimal cutting temperature compound (O.C.T.)</li>
	<li>Label the cryo-mold with a permanent alcohol-resistant marker (mention the orientation of tissue, date and genotype). Prepare a slush of crushed dry ice and 100% ethanol in an ice bucket. Place a glass beaker inside containing isopentane. Wait for 10 min for the isopentane to cool down.</li>
	<li>Meanwhile remove the tissue from the sucrose solution, wipe off excess sucrose around the tissue using laboratory wipes. Transfer the tissue into a pre-labeled cryo-mold with sufficient O.C.T. to cover the tissue. Avoid any air bubbles in the O.C.T.; especially around the tissue.</li>
	<li>Orient the tissue in O.C.T. using featherweight entomology forceps to avoid tissue damage. Start freezing by transferring the cryo-mold into the pre-cooled isopentane bath. Do not splash isopentane into the O.C.T. as it will not freeze properly. Wrap the samples in aluminum foil and store at -80 &#176;C.</li>
</ol>
3. Cryosectioning
<ol>
	<li>Cut 14 &#181;m thin serial sections in series&#8217; of five using a cryostat and collect on SuperFrost&#174; Plus glass slides for immunofluorescence (IF) analysis. Store the slides at -80 &#176;C until utilized.</li>
</ol>
4. Tracer visualization Using the Tyramide Signal Amplification (TSA) Protocol
<ol>
	<li>Prepare the buffers and reagents required for staining. Always freshly prepare the Tris-NaCl-Tween (TNT) buffer on the day of use.
	<ol>
		<li>Prepare TNT buffer using 100 ml of 1 M Tris-HCl pH 7.5, 30 ml of 5 M NaCl and ddH2O up to a final volume of 1 L. Add 500 &#181;l of Tween 20 using a 1 ml pipette. Add the Tween 20 slowly and flush the pipette up and down several times to ensure that all Tween 20 is in solution.</li>
		<li>Prepare Tris-NaCl-Blocking (TNB) buffer using 100 ml of 1 M Tris-HCl pH 7.5, 30 ml of 5 M NaCl and ddH2O up to a final volume of 1 L. Add 5 g Blocking Reagent (provided with the kit) slowly in small increments to the buffer while stirring. Heat the solution gradually to 55 &#176;C with continuous stirring to completely dissolve the Blocking Reagent (this should not take longer than 30-60 min). To achieve homogenous heating use a water bath at 55 &#176;C. The solution will appear milky. Bring the solution to room temperature before using. Aliquot and store at -20 &#176;C for long-term storage.</li>
		<li>Reconstitute biotin tyramide reagent (provided with the kit) in Molecular Biology/HPLC-grade DMSO. Depending upon which kit is used the appropriate amount of DMSO may vary. Store the stock solution at 4 &#176;C. 
		NOTE: Sections from animals heterozygous for the respective Cre and R26 alleles can be used for circuit analysis. Use sections from littermates carrying one Cre or one R26 allele, respectively, as negative controls. Treat negative control slides exactly like slides from double heterozygous animals. In addition, include one control slide from a double heterozygous animal but leave out the TSA reagent (unamplified control).</li>
	</ol>
	</li>
	<li>Using a sharp fresh scalpel remove excess O.C.T. surrounding the tissue sections and mark the boundary around the sections with a PAP pen.</li>
	<li>Dry the slides for 2-3 min at room temperature (RT). Wash the slides at RT 3x with TNT for 5 min each. Incubate the slides at RT for 30 min in 0.3% H2O2 solution in ice-cold methanol to quench endogenous peroxidase activity.</li>
	<li>Wash 3x with PBS for 5 min each at RT. Incubate for 10 min in 0.5% Triton-X 100 in PBS for tissue permeabilization. Wash the slides at RT 3x with TNT for 5 min each.</li>
	<li>Block with TNB (blocking solution, 200-300 &#181;l per slide) for 30 min at RT in a humidified chamber. Make sure all the sections are completely covered by TNB. Do not let the slides dry out in between the steps.</li>
	<li>Drain off excess blocking buffer and apply the primary antiserum recognizing the tracer (1:1,000 goat anti-WGA for BL, 1:15,000 rabbit anti-GFP for GTT) diluted in TNB for 2 hr at RT or overnight at 4 &#176;C. Make sure to use enough volume to completely cover the tissue section (200-300 &#181;l per slide). 
	NOTE: The primary antisera dilutions recommended here may have to be adjusted.</li>
	<li>Wash the slides at RT 3x with TNT for 5 min each with agitation.</li>
	<li>Incubate the sections with biotinylated secondary antiserum recognizing the host of the first antiserum (1:500 horse anti-goat IgG for anti-WGA, 1:5,000 goat anti-rabbit IgG for anti-GFP in TNB for 1 hr at RT).</li>
	<li>Wash 3x with TNT at RT for 5 min each with agitation. Incubate in 1:100 streptavidin (SA)-conjugated horseradish peroxidase (SA-HRP, provided with the kit) in TNB for 30 min at RT in a humidified chamber. Wash the slides at RT 3x with TNT for 5 min each. Meanwhile thaw the TSA.</li>
	<li>Dilute TSA 1:50 in TSA amplification diluent (provided with the kit; use the TSA kit for the visualization of BL and the TSA+ kit for GTT). Incubate the tissue sections in diluted TSA for precisely 10 min at RT. Dilute TSA just before use, ideally during the previous washing step, do not use any remaining diluted TSA reagent for future experiments; always prepare fresh.</li>
	<li>Wash the slides at RT 3x with TNT for 5 min each with agitation. Incubate with 1:500 Alexa Fluor&#174; 546 streptavidin conjugate in TNB for 30 min at RT. Wash the slides at RT 3x with TNT for 5 min each with agitation.</li>
	<li>Incubate for 10 min in 5% bisbenzimide solution in PBS at RT for nuclear staining. Wash the slides at RT 3x with TNT for 5 min each with agitation. Mount with Fluromount G and perform epifluorescence or confocal microscopy. 
	Troubleshooting: Efficacy of tracer transfer will depend on the expression level of the ROSA26 locus in the producing cells (determined by the Cre driver line used) and on the neural circuit analyzed (e.g. number of neurons producing the tracer, convergence etc.). Therefore the primary antisera dilutions recommended here may have to be adjusted to prevent low or excess signal. Background should always be analyzed using appropriate control slides (see above).</li>
</ol>
5. &#964;lacZ Staining for Embryonic Sections
<ol>
	<li>Prepare the buffers and reagents required for staining. Always freshly prepare the LacZ buffer C before use.
	<ol>
		<li>Prepare 5-bromo-4-chloro-3-indolyl-&#946;-D-galactopyranoside (X-gal) stock solution (40 mg/ml) by adding 40 g of X-gal to 1 ml 70% dimethylformamide (DMF). Cover with aluminium foil and store at -20 &#176;C for long-term storage.</li>
		<li>Prepare nitro blue tetrazolium (NBT) stock solution (50 mg/ml) by adding 50 g of NBT to 1 ml 70% DMF. Cover with aluminium foil and store at -20 &#176;C for long-term storage.</li>
		<li>Prepare LacZ fixative using 80 &#181;l of 25% glutaraldehyde, 5 ml of 4% PFA, 20 &#181;l of 1 M MgCl2, 100 &#181;l of 500 mM EGTA, 1 ml of 1 M phosphate buffer pH 7.4 and ddH2O up to a final volume of 10 ml.</li>
		<li>Prepare LacZ buffer A using 1 ml of 1 M MgCl2, 5 ml of 500 mM EGTA, 50 ml of 1 M phosphate buffer pH 7.4 and ddH2O up to a final volume of 500 ml.</li>
		<li>Prepare LacZ buffer B using 1 ml of 1 M MgCl2, 5 ml of 500 mM EGTA, 5 ml of 1% sodium deoxycholate, 100 ml of 10% NP-40, 50 ml of 1 M phosphate buffer pH 7.4 and ddH2O up to a final volume of 500 ml.</li>
		<li>Prepare LacZ buffer C by adding 10 &#181;l of NBT stock solution and 12.5 &#181;l of X-gal stock solution to 1 ml of LacZ buffer B. Make fresh just before use and cover with aluminium foil.</li>
	</ol>
	</li>
	<li>Dry the slides at RT at least for 10-15 min (meanwhile prepare the slides with a PAP pen).</li>
	<li>Fix the section with LacZ fixative for 2 min at room temperature in a humidified chamber inside a chemical fume-hood. Wash 3 times with PBS supplemented with 2 mM MgCl2.</li>
	<li>Rinse the slides with LacZ buffer A. Wash the slides with LacZ buffer A 3 times for 10 min each at RT. Wash 2 times with LacZ buffer B for 5 min each. Incubate the slides in LacZ buffer C at 30-37 &#176;C for 6 hr or overnight in a humidified chamber. 
	NOTE: Add sufficient volume of LacZ buffer C; slides should not dry out during incubation.</li>
	<li>After appropriate incubation wash the slides with PBS supplemented with 2 mM MgCl2. Rinse sections briefly with ddH2O. Coverslip with Mayer&#8217;s glycine gelatin. Suitable for a light microscope equipped with a differential interference contrast (DIC) imaging set-up.</li>
</ol>
6. Gender and Tracer Genotyping
<ol>
	<li>Prepare tail lysis buffer using 1 ml of 1 M Tris-HCl pH 8.5, 100 &#181;l of 500 mM EGTA, 200 &#181;l of 10% SDS, 400 &#181;l of 5 M NaCl and ddH2O to make up a final volume of 10 ml.</li>
	<li>Add 500 &#181;l of tail lysis buffer to each Eppendorf tube containing a tail biopsy. Add proteinase K to a final concentration of 100 mg/ml. Incubate overnight on a shaker at 55 &#176;C.</li>
	<li>Centrifuge samples at 17,000 x g for 10 min at RT. Transfer the supernatant to a new centrifuge tube. Add 500 &#181;l of isopropanol to the supernatant. Mix for at least 5 min by inverting the tubes up and down.</li>
	<li>Spin in centrifuge at 17,000 x g for 5 min at RT. Pour off the supernatant. Add 200 &#181;l of 70% ethanol. Shake for at least 5 min.</li>
	<li>Centrifuge at 17,000 x g for 5 min. Remove supernatant with a Pipetman (do not pour off). Dry the pellet in a warm chamber (37 &#176;C) for 15-30 min.</li>
	<li>Resuspend pellet in 100 &#181;l of low TE pH 8.0 or water using wide-bore filtered tips. Put in 55 &#176;C for 1 hr and at 37 &#176;C overnight for proper dissolving. Store at 4 &#176;C.</li>
	<li>Use 1 &#181;l of DNA for PCR reaction (50 &#956;l reaction mixture containing 1 &#956;l of tail DNA, 2.5 &#956;l DMSO, 10 pM of each primer, 25 &#956;M of each dNTP, and 1M betaine). PCR primers for genotyping and gender identification are listed in the Materials table.</li>
</ol>

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+half+century+of+experimental+neuroanatomical+tracing.">A half century of experimental neuroanatomical tracing.</a> J. Chem. 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No conflict of interest declared.

Expressing transsynaptic tracers as transgenes to trace the neural circuits of genetically defined neuronal populations has several advantages compared to the stereotaxic injection of tracers or neurotopic viruses. First, the tracer is produced as an endogenous protein and therefore does not elicit any immune response and a selective neural pathway can be analyzed in different animals with high reproducibility. Second, because this is a non-invasive method it can be utilized to trace the circuits from neurons not easily accessible for stereotaxic injections, for example in utero. Limitations include a generally low efficacy in transsynaptic transfer, which may result in difficulties in detecting the tracer. Furthermore, the tracer molecule gets diluted at every synapse. In contrast, neurotropic viruses replicate and viral replication in turn then leads to signal amplification after crossing the synapse. However, viral replication is also a major limitation of the use of neurotropic viruses due to intrinsic viral cytotoxicity. Some features of genetic transsynaptic tracers and genetically modified neurotropic viruses are compared in Table 1.
The R26-BIZ and R26-GTT alleles complement each other and facilitate the visualization of the neural circuits of a genetically identified neuronal population upon Cre-mediated recombination. The R26 promoter is well characterized and mediates ubiquitous expression with modest strength11,12. Using highly sensitive methods for tracer detection such as the tyramide signal amplification (TSA) system allows the identification of even minute amounts of tracer molecules transferred across synapses. Another major advantage of using the R26 promoter to drive tracer expression is that the expressing neurons continuously synthesize the tracers throughout their life. This makes it possible to analyze circuit maturation for prolonged time periods5. In contrast, this is not possible when using neurotropic viruses due to their cytotoxicity. Importantly, our binary genetic system and the use of the R26 promoter uncouples tracer production from potentially heavily regulated endogenous promoters driving Cre expression (in this case, the Kiss1 promoter), which may be regulated by a variety of genetic and epigenetic factors13. Hence transsynaptic transfer of BL and GTT reflects actual synaptic communication between two neuronal populations.
Cre-mediated recombination is an irreversible event and therefore it converts developmentally transient gene expression into stable transgene expression. The use of an inducible Cre system14 can potentially unmask a developmental effect due to transient Cre expression.
In conclusion, the two novel R26-BIZ and R26-GTT mouse strains can be used to analyze local and long range neural circuits that might be composed of different classes and types of neurons originating from any brain region or even from the spinal cord in a Cre-dependent manner.

Anatomical path tracing is one of the most commonly utilized tools to decipher the relationship between brain and behavior1. Advancement in neural circuit tracing technologies has bestowed neuroscientists with the ability to trace neural circuits from genetically identified neuron populations in mice2. In spite of these technical advancements it remains challenging to unravel the formation of neural circuits especially during embryonic maturation. This is because most of the tracing methods developed to date are based upon stereotaxic injection of transsynaptic tracers or genetically modified neurotropic viruses (Figure 1)2,3. While these techniques achieve spatial and temporal resolution of connectivity, several inherent limitations such as technically challenging tracer injections into the developing brain, the reproducibility of the site of injection, potential inflammation at the injection site and most importantly cytotoxicity caused by neurotropic viruses limit their use4.
An alternative method is to express the transsynaptic tracers as transgenes in genetically altered mice. We have recently modified this technique and developed a binary genetic transsynaptic tracing system to map the neural circuits of any genetically identified neuronal population5. Our experimental strategy is based on two new knock-in mouse strains, which express either the bidirectional tracer barley lectin (BL)6 or the retrograde tracer Tetanus Toxin fragment C fused to GFP (GTT)7 from the ROSA26 locus after Cre-mediated recombination. Here we used these mouse strains to selectively express BL and GTT in neurons that produce kisspeptin, a neuropeptide that is implicated in regulating the maturation of the reproductive axis8,9. We demonstrate that this technique is suitable to visualize the development and maturation of kisspeptin neural circuitry during embryonic development of the female mouse brain5.
Breeding strategy
The R26-BL-IRES-&#964;lacZ (BIZ) and the R26-GFP-TTC (GTT) tracer lines are knock-in strains5 that carry recombinant ROSA26 alleles. The R26-BIZ and the R26-GTT alleles are transcriptionally silent due to the presence of a strong transcriptional stop signal, which is flanked by two loxP sites5. Expression of the BIZ and GTT transgene is activated by Cre-mediated removal of the transcriptional stop signal. The R26-BIZ and R26-GTT alleles can be used independently by simply crossing with a Cre driver line. For analysis animals heterozygous for the respective Cre and R26 alleles can be used. Littermates carrying one Cre or one R26 allele, respectively, should be used as controls. Alternatively, it is also possible to generate triple knock-in animals carrying the Cre, R26-BIZ and R26-GTT alleles, however this will require one additional cross.

<table><tbody><tr><td>Bisbenzimide (Hoechst 33258 dye)</td><td>Sigma</td><td>14530-100MG</td><td></td></tr><tr><td>Ethanol</td><td>Sigma</td><td>32205-1L</td><td></td></tr><tr><td>Cryo mold (Peel-a-way)</td><td>Polyscience Inc.</td><td>18646A-1</td><td>22 x 22 x 20mm</td></tr><tr><td>DMSO</td><td>Sigma</td><td>D8418-100ML</td><td></td></tr><tr><td>Dimethyl Formamide (DMF)</td><td>VWR Chemicals</td><td>23470,293</td><td></td></tr><tr><td>EGTA</td><td>ROTH</td><td>3054.3</td><td></td></tr><tr><td>Fluoromount G</td><td>Southern Biotech</td><td>0100-01</td><td></td></tr><tr><td>Glutaraldehyde</td><td>Sigma</td><td>G5882-50ML</td><td></td></tr><tr><td>Hydrogen peroxide</td><td>Sigma</td><td>34988-7</td><td></td></tr><tr><td>Isopentane (Methyl 2-butane)</td><td>Sigma</td><td>M32631-2.5L</td><td></td></tr><tr><td>Kaiser&#039;s glycine gelatin</td><td>Merck</td><td>1092420100</td><td></td></tr><tr><td>Methanol</td><td>Sigma</td><td>494437-1L</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M2670-100G</td><td></td></tr><tr><td>NaCl</td><td>ROTH</td><td>HN00.2</td><td></td></tr><tr><td>NBT</td><td>Sigma</td><td>298-83-9</td><td></td></tr><tr><td>Nonidet P40 substitute</td><td>Fluka</td><td>743.85</td><td></td></tr><tr><td>OCT</td><td>Leica</td><td>14020108926</td><td></td></tr><tr><td>PAP pen</td><td>Dako</td><td>S2002</td><td></td></tr><tr><td>Parafarmaldehyde</td><td>Sigma</td><td>P6148-1KG</td><td></td></tr><tr><td>Sodium deoxycholate</td><td>Sigma</td><td>D6750-25G</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S7903-1KG</td><td></td></tr><tr><td>Superfrost slides</td><td>Thermo Scientific</td><td>FT4981GLPLUS</td><td></td></tr><tr><td>TSA kit</td><td>PerkinElmer</td><td>NEL700</td><td></td></tr><tr><td>TSA plus kit</td><td>PerkinElmer</td><td>NEL749A001KT</td><td></td></tr><tr><td>Tris</td><td>ROTH</td><td>AE15.2</td><td></td></tr><tr><td>Triton-X 100</td><td>ROTH</td><td>3051.2</td><td></td></tr><tr><td>Tween 20</td><td>ROTH</td><td>9127.1</td><td></td></tr><tr><td>X-gal</td><td>ROTH</td><td>2315.1</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>na</td><td></td></tr><tr><td>Light microscope equipped with DIC imaging</td><td>Zeiss</td><td>Axioskop2 equipped with Axio Vision software</td><td></td></tr><tr><td>Fluroscence microscope</td><td>Zeiss</td><td>Axioskop2 equipped with Axio Vision software</td><td></td></tr><tr><td>Photoshop</td><td>Adobe</td><td>PS6</td><td></td></tr><tr><td>Goat anti-WGA (recognizes BL)</td><td>Vector Laboatories</td><td>AS-2024</td><td></td></tr><tr><td>Biotinylayted horse anti-goat IgG</td><td>Vector Laboatories</td><td>BA-9500</td><td></td></tr><tr><td>Biotinylated goat anti-rabbit IgG</td><td>Vector Laboatories</td><td>BA-1000</td><td></td></tr><tr><td>Rabbit anti-GFP (recognizes GTT)</td><td>Invitrogen</td><td>A11122</td><td></td></tr><tr><td>Rabbit anti-GnRH</td><td>Affinity Bio Reagent</td><td>PA1-121</td><td></td></tr><tr><td>Dylight488-donkey anti-rabbit IgG</td><td>Thermo Scientific</td><td>SA5-10038</td><td></td></tr><tr><td>SA-Alexa Fluor 546</td><td>Life Technologies</td><td>S-11225</td><td></td></tr><tr><td colspan="4">Primers</td></tr><tr><td>BL Fwd (for BIZ genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>ATGAAGATGATGAGCACCAGGGC</td></tr><tr><td>BL Rev (for BIZ genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>AGCCCTCGCCGCAGAACTC</td></tr><tr><td>Cre Fwd (for Cre genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>GTCGATGCAACGAGTGATGAGGTTCG</td></tr><tr><td>Cre Rev (for Cre genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>CCAGGCTAAGTGCCTTCTCTACACCTGC</td></tr><tr><td>TTC Fwd (for GTT genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>AGCAAGGGCGAGGAGCTGTT</td></tr><tr><td>TTC Rev (for GTT genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>GTCTTGTAGTTGCCGTCGTCCTTGAA</td></tr><tr><td>XY Fwd (for gender genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>TGAAGCTTTTGGCTTTGA</td></tr><tr><td>XY Rev (for gender genotyping)</td><td>Eurofins MWG Operon</td><td></td><td>CCGCTGCCAAATTCTTTG</td></tr><tr><td>ROSA26 Fwd</td><td>Eurofins MWG Operon</td><td></td><td>CGAAGTCGCTCTGAGTTGTTATC</td></tr><tr><td>ROSA26 Rev</td><td>Eurofins MWG Operon</td><td></td><td>GCAGATGGAGCGGGAGAAAT</td></tr><tr><td>SA Rev</td><td>Eurofins MWG Operon</td><td></td><td>CGAAGTCGCTCTGAGTTGTTATC</td></tr></tbody></table>

conditional genetic transsynaptic tracing in the embryonic mouse brain

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits during embryonic maturation of the brain however is technically challenging because most transsynaptic tracing methods developed to date depend on stereotaxic tracer injection. To overcome this problem, we developed a binary genetic strategy for conditional genetic transsynaptic tracing in the mouse brain. Towards this end we generated two complementary knock-in mouse strains to selectively express the bidirectional transsynaptic tracer barley lectin (BL) and the retrograde transsynaptic tracer Tetanus Toxin fragment C from the ROSA26 locus after Cre-mediated recombination. Cell-specific tracer production in these mice is genetically encoded and does not depend on mechanical tracer injection. Therefore our experimental approach is suitable to study neural circuit formation in the embryonic murine brain. Furthermore, because tracer transfer across synapses depends on synaptic activity, these mouse strains can be used to analyze the communication between genetically defined neuronal populations during brain development at a single cell resolution. Here we provide a detailed protocol for transsynaptic tracing in mouse embryos using the novel recombinant ROSA26 alleles. We have utilized this experimental technique in order to delineate the neural circuitry underlying maturation of the reproductive axis in the developing female mouse brain.

This section shows representative results that can be obtained working with the R26-BIZ (BL-IRES-&#964;lacZ) and the R26-GTT (GFP-TTC) alleles. Here we use the R26-BIZ and the R26-GTT alleles to analyze the maturation of the neural circuits regulating the reproductive axis. Reproduction in vertebrates is centrally controlled by a small subset of neurons in the hypothalamus, which secrete gonadotropin-releasing hormone (GnRH). Kisspeptin, a potent activator of GnRH neurons, has been implicated in regulating the activity of GnRH neurons, however when the neural circuits between GnRH and kisspeptin neurons are established in the developing female mouse brain was not known5. First we show that expression of the BIZ and GTT transgenes are specifically activated by Cre-dependent recombination in embryonic kisspeptin neurons in the arcuate nucleus (ARC) (Figure 2) using a kisspeptin-specific Cre driver line10. We then demonstrate that kisspeptin neurons in the ARC already communicate with a specific subset of GnRH neurons in utero (Figure 3A, B). Furthermore, we show that ARC kisspeptin neurons are upstream of GnRH neurons (Figure 3A, C). Taken together our findings indicate that the neural circuits between kisspeptin and GnRH neurons are fully established and operative in the female mouse brain before birth.
<img alt="Figure 1" src="/files/ftp_upload/52487/52487fig1highres.jpg" /> 
Figure 1: Transsynaptic transfer of tracers using conventional or genetic approaches. In the conventional approach, a transsynaptic tracer is taken up by all the neurons at the injection site thus potentially labeling non-specific unrelated pathways. In the genetic approach, the tracer is selectively expressed by genetically defined neurons thus specifically visualizing neurons connected to the tracer-expressing cells. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52487/52487fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52487/52487fig2highres.jpg" width="600" /> 
Figure 2: Faithful activation of the BIZ and GTT transgene in embryonic ARC kisspeptin neurons. (A) Breeding strategy to activate BIZ and GTT expression in kisspeptin neurons. We bred R26-BIZ and R26-GTT mice with Kisspeptin-IRES-Cre (KissIC) mice10, in which Cre-recombinase is expressed under the control of the Kiss1 promoter. (C-E) &#946;-gal enzymatic activity identifies the tracer-producing cells and is restricted to the ARC in the brain of a female KissIC/R26-BIZ embryo at embryonic day (E) 18.5. (F-G) Double immunofluorescence for kisspeptin (green) and BL (red) (F) or GTT (red) (G) demonstrates faithful activation of BL or GTT expression by Cre-mediated recombination in kisspeptin neurons in KissIC/R26-BIZ or KissIC/R26-GTT female mouse embryos, respectively. Scale bars (C) 500 &#181;m, (D) 200 &#181;m, (F, G) 50 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52487/52487fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52487/52487fig3highres.jpg" width="600" /> 
Figure 3: GnRH neurons are synaptically connected to and downstream of kisspeptin neurons. (A) Combinatorial genetic bidirectional transsynaptic tracing. While &#964;lacZ is confined to the Cre-expressing kisspeptin neurons, BL is transsynaptically transferred to upstream (presynaptic) and downstream (postsynaptic) neurons in KissIC/R26-BIZ mice. In contrast, GFP-TTC is transsynaptically transferred to presynaptic, but not postsynaptic neurons in KissIC/R26-GTT mice. Postsynaptic neurons have &#964;lacZ-positive axon fibers in their close vicinity. (B, C) Double immunofluorescence for BL (red; B) or GTT (red; C) and GnRH (green) on a sagittal section through the whole head of a female KissIC/R26-BIZ or KissIC/R26-GTT embryo at E18.5. (B) Note that some, but not all GnRH neurons contain BL. These data demonstrate that a subset of embryonic GnRH neurons is synaptically connected to ARC kisspeptin neurons. (C) Note that none of the GnRH neurons contain GTT. Exclusive transsynaptic transfer of BL but not GTT to GnRH neurons demonstrates that GnRH neurons are downstream of kisspeptin neurons. Scale bars (B, C) 50 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/52487/52487fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Features</td>
			<td>Transsynaptic tracers</td>
			<td>Neurotropic viruses</td>
		</tr>
		<tr>
			<td>Mode of application</td>
			<td>Expressed as a transgene</td>
			<td>Mechanically injected</td>
		</tr>
		<tr>
			<td>Direction of spread</td>
			<td>Anterograde, retrograde &#38; bidirectional</td>
			<td>Retrograde &#38; anterograde</td>
		</tr>
		<tr>
			<td>Synaptic efficacy</td>
			<td>Multisynaptic</td>
			<td>Multisynaptic or monosynaptic (retrograde only)</td>
		</tr>
		<tr>
			<td>Signal strength</td>
			<td>Weak, decreases at every synapse</td>
			<td>Strong, signal amplification at every synapse</td>
		</tr>
		<tr>
			<td>Immune response&#160;</td>
			<td>None; expressed as endogenous protein</td>
			<td>Strong immune response, lethal</td>
		</tr>
		<tr>
			<td>Spatial and temporal resolution</td>
			<td>Depends on promoter of choice</td>
			<td>Highly selective due to mechanical injections at individual sites</td>
		</tr>
	</tbody>
</table>
Table 1: Comparison of features of genetic transsynaptic tracers and genetically modified neurotropic viruses

Watch this Scientific Journal Video about Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain at JoVE.com

Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain

Capitalizing on a binary genetic strategy we provide a detailed protocol for neural circuit tracing in mice that express complementary transsynaptic tracers after Cre-mediated recombination. Because cell-specific tracer production is genetically encoded, our experimental approach is suitable to study the formation and maturation of neural circuitry during murine embryonic brain development at a single cell resolution.

Anatomical path tracing is of pivotal importance to decipher the relationship between brain and behavior. Unraveling the formation of neural circuits ...

conditional-genetic-transsynaptic-tracing-in-the-embryonic-mouse-brain

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

18.6K Views.  University of Saarland School of Medicine. Capitalizing on a binary genetic strategy we provide a detailed protocol for neural circuit tracing in mice that express complementary transsynaptic tracers after Cre-mediated recombination. Because cell-specific tracer production is genetically encoded, our experimental approach is suitable to study the formation and maturation of neural circuitry during murine embryonic brain development at a single cell resolution. 

Video: Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain

Mosaic Analysis of Gene Function in Postnatal Mouse Brain Development by Using Virus-based Cre Recombination

Normal brain function relies not only on embryonic development when major neuronal pathways are established, but also on postnatal 
development when neural circuits are matured and refined. Misregulation at this stage may lead to neurological and psychiatric disorders such as autism 
and schizophrenia1,2. Many genes have been studied in the prenatal brain and found crucial to many developmental processes3-5. However, their 
function in the postnatal brain is largely unknown, partly because their deletion in mice often leads to lethality during neonatal development, and partly because their requirement in early development hampers the postnatal analysis. To overcome these obstacles, floxed alleles of these genes are currently being generated in mice 6. When combined with transgenic alleles that express Cre recombinase in specific cell types, conditional deletion can be achieved to study gene function in the postnatal brain. However, this method requires additional alleles and extra time (3-6 months) to generate the mice with appropriate genotypes, thereby limiting the expansion of the genetic analysis to a large scale in the mouse brain.
Here we demonstrate a complementary approach that uses virally-expressed Cre to study these floxed alleles rapidly and 
systematically in postnatal brain development. By injecting recombinant adeno-associated viruses (rAAVs)7,8 encoding Cre into the neonatal brain, 
we are able to delete the gene of interest in different regions of the brain. By controlling the viral titer and coexpressing a fluorescent 
protein marker, we can simultaneously achieve mosaic gene inactivation and sparse neuronal labeling. This method bypasses the requirement of 
many genes in early development, and allows us to study their cell autonomous function in many critical processes in postnatal brain development,
 including axonal and dendritic growth, branching, and tiling, as well as synapse formation and refinement. This method has been used successfully 
in our own lab (unpublished results) and others8,9, and can be extended to other viruses, such as lentivirus 9, as well as to the expression of 
shRNA or dominant active proteins 10. Furthermore, by combining this technique with electrophysiology as well as recently-developed optical 
imaging tools 11, this method provides a new strategy to study how genetic pathways influence neural circuit development and function in mice 
and rats.

An in vivo method to test gene function in postnatal brain is described. Recombinant AAVs expressing Cre and/or a fluorescent protein are injected into neonatal mouse brain. Mosaic gene inactivation and sparse neuronal labeling are achieved, allowing rapid analysis of gene function in processes critical to neural circuit development.

Normal brain function relies not only on embryonic development when major neuronal pathways are established, but also on postnatal 
development when ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Viral Tracing of Genetically Defined Neural Circuitry

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus (RABV), and more recently vesicular stomatitis virus (VSV), for studying circuitry, is becoming increasingly popular. These higher throughput methods use viruses that transmit between neurons in either the anterograde or retrograde direction.
Recently, a modified RABV for monosynaptic retrograde tracing was developed. (Figure 1A). In this method, the glycoprotein (G) gene is deleted from the viral genome, and resupplied only in targeted neurons. Infection specificity is achieved by substituting a chimeric G, composed of the extracellular domain of the ASLV-A glycoprotein and the cytoplasmic domain of the RABV-G (A/RG), for the normal RABV-G1. This chimeric G specifically infects cells expressing the TVA receptor1. The gene encoding TVA can been delivered by various methods2-8. Following RABV-G infection of a TVA-expressing neuron, the RABV can transmit to other, synaptically connected neurons in a retrograde direction by nature of its own G which was co-delivered with the TVA receptor. This technique labels a relatively large number of inputs (5-10%)2 onto a defined cell type, providing a sampling of all of the inputs onto a defined starter cell type.
We recently modified this technique to use VSV as a transsynaptic tracer9. VSV has several advantages, including the rapidity of gene expression. Here we detail a new viral tracing system using VSV useful for probing microcircuitry with increased resolution. While the original published strategies by Wickersham et al.4 and Beier et al.9 permit labeling of any neurons that project onto initially-infected TVA-expressing-cells, here VSV was engineered to transmit only to TVA-expressing cells (Figure 1B). The virus is first pseudotyped with RABV-G to permit infection of neurons downstream of TVA-expressing neurons. After infecting this first population of cells, the virus released can only infect TVA-expressing cells. Because the transsynaptic viral spread is limited to TVA-expressing cells, presence of absence of connectivity from defined cell types can be explored with high resolution. An experimental flow chart of these experiments is shown in Figure 2. Here we show a model circuit, that of direction-selectivity in the mouse retina. We examine the connectivity of starburst amacrine cells (SACs) to retinal ganglion cells (RGCs).

A method of tracing synaptically connected neurons is described. We use TVA specificity of an upstream cell to probe whether a cell population of interest receives synaptic input from genetically defined cell types.

Classical methods for studying neuronal circuits are fairly low throughput. Transsynaptic viruses, particularly the pseudorabies (PRV) and rabies virus ...

Electroporation of the Hindbrain to Trace Axonal Trajectories and Synaptic Targets in the Chick Embryo

Electroporation of the chick embryonic neural tube has many advantages such as being quick and efficient for the expression of foreign genes into neuronal cells. In this manuscript we provide a method that demonstrates uniquely how to electroporate DNA into the avian hindbrain at E2.75 in order to specifically label a subset of neuronal progenitors, and how to follow their axonal projections and synaptic targets at much advanced stages of development, up to E14.5. We have utilized novel genetic tools including specific enhancer elements, Cre/Lox - based plasmids and the PiggyBac-mediated DNA transposition system to drive GFP expression in a subtype of hindbrain cells (the dorsal most subgroup of interneurons, dA1). Axonal trajectories and targets of dA1 axons are followed at early and late embryonic stages at various brainstem regions. This strategy contributes advanced techniques for targeting cells of interest in the embryonic hindbrain and for tracing circuit formation at multiple stages of development.

How neuronal networks are established in the embryonic brain is a fundamental question in developmental neurobiology. Here we combined an electroporation technique with novel genetic tools, such as Cre/Lox&#x2013;plasmids and PiggyBac-mediated DNA transposition system in the avian hindbrain to label dorsal interneurons and track their axonal projections and synaptic targets at various developmental stages.

Electroporation of  the chick embryonic neural tube has many advantages such as being quick and efficient  for the expression of foreign genes into ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

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

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

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

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

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Laser-guided Neuronal Tracing In Brain Explants

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake through electroporation in brain explants with targeted laser illumination and matching filter goggles during the procedure. The described technique of in vitro electroporation by itself yields relatively good visual control for targetting certain areas of the brain. By combining it with laser excitation of fluorescent genetic markers and their read-out through band-passing filter goggles, which can pick up the emissions of the genetically labeled cells/nuclei and the fluorescent tracing dye, a researcher can substantially increase the accuracy of injections by finding the area of interest and controlling for the dye-spread/uptake in the injection area much more efficiently. In addition, the laser illumination technique allows to study the functionality of a given neurocircuit by providing information about the type of neurons projecting to a certain area in cases where the GFP expression is linked to the type of transmitter expressed by a subpopulation of neurons.

We describe a technique to label neurons and their processes via anterograde or retrograde tracer injections into brain nuclei using an in vitro preparation. We modified an existing method of in vitro tracer electroporation by taking advantage of fluorescently labeled mouse mutants and basic optical equipment in order to increase labeling accuracy.

We present a technique which combines an in vitro tracer injection protocol, which uses a series of electrical and pressure pulses to increase dye uptake ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

Conditional Genetic Transsynaptic Tracing in the Embryonic Mouse Brain

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Chapters in this video

Mosaic Analysis of Gene Function in Postnatal Mouse Brain Development by Using Virus-based Cre Recombination

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

Viral Tracing of Genetically Defined Neural Circuitry

Electroporation of the Hindbrain to Trace Axonal Trajectories and Synaptic Targets in the Chick Embryo

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Laser-guided Neuronal Tracing In Brain Explants

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation