This work is funded by CBS Bridging Grant awarded to OOM. OOM, PAA, and AS designed the study and performed the experiments. AS and PAA analyzed the results. OOM and PAA prepared the manuscript.&#160;We thank Dr. Karl Disseroth (Stanford University) for making the AAV available for our use.

All experimental and animal handling procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of the Louisiana State University School of Veterinary Medicine.
1. Experimental animal
<ol>
	<li>Use 5&#8211;6 weeks old mice.</li>
	<li>House 3&#8211;5 animals per cage under standard conditions of 12 h alternating light and dark cycle. Food and water should be provided ad libitum.</li>
</ol>
2. Craniotomy and animal preparation
NOTE: This section describes pre- and peri-operative procedures for mouse craniotomy. Use standard stereotactic apparatus and appropriate coordinates (Anteroposterior: AP, Mediolateral: ML, and Dorsoventral: DV). Refer to a mouse brain atlas to determine the coordinates for the brain region of interest.
<ol>
	<li>Anesthetize mice by intraperitoneal ketamine (100 mg/kg)/xylazine (10 mg/kg) cocktail injection. Perform a toe pinch test to ensure the absence of sensation before the commencement of surgery. 
	NOTE: Alternatively, Isoflurane anesthesia with an appropriate nose chamber can also be used for this step.</li>
	<li>Gently fix the head of the animal on the stereotaxic apparatus. 
	NOTE: It is important to handle animals with caution during this process. Also, check the breathing rate and other vitals before proceeding with surgery. Allow the animal to rest for ~7 min to reduce stress.
	<ol>
		<li>Place a heating pad on the stereotaxic frame such that the body of the animal is lying on it. This will help maintain the body temperature and keep the animal warm though out the procedure. Retain the heating pad for postoperative care and recovery.</li>
	</ol>
	</li>
	<li>Use a clipper to remove hair over the scalp and clean the skin with an iodine solution.
	<ol>
		<li>Apply topical lidocaine to block sensation on the scalp.</li>
		<li>Use a scalpel to make a midline scalp incision extending from the frontal to the occipital region.</li>
		<li>Clean the incision area with iodine solution, then expose the calvaria. 
		NOTE: A small amount of 3% hydrogen peroxide solution can be applied to remove periosteum from the calvaria. This will increase the visibility of landmarks (bregma and lambda) and sutures.</li>
		<li>Using a mouse brain stereotactic atlas, determine AP and ML coordinates relative to the bregma.</li>
		<li>For VTA injection, position an ultrafine blunt-point needle syringe (e.g., Neuros syringe) at coordinates AP: -3.08 mm/ML: 0.5 mm relative to the bregma.</li>
		<li>Use a drilling tool to bore a 1 mm hole (craniotomy) in the skull at the marked AP/ML coordinate. 
		NOTE: This step requires a significant level of caution. Minimal pressure should be applied while drilling to prevent the drill bit from crushing the brain tissue. In the current study, drill bit was 0.8 mm and drill speed was set to 15,000 rpm. 
		CAUTION: If hydrogen peroxide was used in cleaning the incision or calvaria, ensure total removal of solution or allow complete dryness before drilling.</li>
	</ol>
	</li>
</ol>
3. AAV cocktail injection
NOTE: This section describes the process for AAV injection into the VTA of adult C57BL/6 mice (23&#8211;27 g). The described method can be used for AAV injection into any brain region using standard stereotactic apparatus and appropriate coordinates.&#160;To demonstrate this protocol, eYFP and hChR2 were expressed in VTA glutamate neurons using AAV5-mediated transfection under a CaMKII&#945; promoter (Figure 1). Cre-lox recombination can also be used for this step.
<ol>
	<li>Mount a syringe with ultrafine blunt-point needle (32 G; 5 &#181;L capacity with 100 nL accuracy) on an injector. Fix the syringe holder on a micromanipulator. 
	NOTE: For this procedure, a manual syringe holder, with 1.6 nL calibration, was used. An automated injector can also be used.</li>
	<li>Fill the syringe with double distilled water to clean and test the flow of fluid.</li>
	<li>Thaw aliquots of AAV (serotype 5) cocktail on ice. AAVs are best stored at -80 &#176;C to maintain viral titer for good expression.</li>
	<li>Fill the mounted syringe with 1,000 nL of AAV solution (10 mM in phosphate-buffered saline (PBS), pH 7.4). Dispense 10 nL of the AAV solution to confirm the flow of the liquid before lowering the needle into an injection site.</li>
	<li>Use a micromanipulator to lower the&#160;needle stereotactically into the desired depth (DV coordinate). After lowering the needle to the desired depth, allow the syringe to remain in place for 10 min before injection. 
	NOTE: For this procedure, the needle was lowered into the VTA at a depth (DV) of 4.5 mm from the pial surface of the brain. For other brain areas, use the appropriate dorsoventral coordinate (refer to the mouse brain atlas).</li>
	<li>Inject 600&#8211;800 nL of AAV into the VTA. Injection volume may be adjusted depending on the size of the target site.
	<ol>
		<li>Deliver the AAV solution at the rate of 60 nL/min (3 min interval).</li>
		<li>To express eYFP and hChR2 in VTA glutamate neurons and projections, inject AAV-CaMKII&#945;-ChR2-eYFP. 
		NOTE: Cre-lox recombination method can also be used depending on the goal of the experiment.</li>
		<li>Allow the needle to remain in place for 15 min after AAV injection. This will&#160;reduce diffusion and backflow.</li>
	</ol>
	</li>
	<li>Withdraw the syringe needle, and then suture the incision to close the wound.</li>
	<li>Administer antibiotics and analgesia as a part of post-operative care. Place the mouse&#160;on a warm padded platforms and monitor the animal until awake. 
	NOTE: After 3 weeks of injection, AAV expression can be observed by fluorescence detection of reporter protein (eYFP) in mouse brain sections. Also, photostimulation experiments can be performed after 3 weeks (Figure 2).</li>
</ol>
4. Set up for in vivo neural recordings with optogenetics
NOTE: This section describes the steps for acute neural recording with the optogenetic tracing of a brain circuit (VTA glutamate neuron projection to the CA1). If necessary, check the system (amplifier and connections) for electrical noise and grounding issues before commencing this step. Performing this step in a Faraday cage can help eliminate electrical noise in the recording.
<ol>
	<li>At 3 weeks after AAV injection, anesthetize mice by intraperitoneal urethane injection (0.2 mg/kg). 
	CAUTION: Urethane is carcinogenic and must be handled cautiously while wearing protective gear. Also, administering urethane anesthesia at this concentration is non-survival.</li>
	<li>Affix the head of the animal on a stereotactic frame as described previously (step 2.2, Figure 3A). Perform a craniotomy to expose the dura (Figure 3A). Use a drilling tool (bit size: 0.8 mm, speed: 15,000 rpm) to remove part of the parietal bone.
	<ol>
		<li>The craniotomy should be about 3 mm x 4mm wide. Apply drops of artificial cerebrospinal fluid (aCSF) over this area to prevent dryness.</li>
	</ol>
	</li>
	<li>Under a dissection (digital) microscope, use a bent needle tip (27 G) to excise the exposed dura. Be careful not to pull apart the delicate pial covering and cortical tissues in this area.</li>
	<li>Bore a hole in the occipital&#160;bone (bit size: 0.8 mm, speed: 10,000 rpm) to hold the ground screw (pan head Phillips screw; M0.6 x 2.0 mm).</li>
	<li>Connect a stainless-steel ground wire. 
	NOTE: Other types of wires can also be used (Figure 3b).</li>
	<li>For combined photostimulation (VTA) and neural recording (CA1), use a multi-rail stereotaxic apparatus fitted with ultra-fine (10 &#181;m or 1 &#181;m) micromanipulators. Mount the optical fiber and recording neural probe on 10 &#956;m-range and 1 &#956;m-range micromanipulator, respectively.
	<ol>
		<li>If necessary, use a head stage-probe adapter (Figure 3c). 
		NOTE: In the current protocol, a 32-channel head stage was fitted with an adapter to hold a 4-channel recording electrode (Figure 3d). The stainless-steel ground wire was soldered to the ground connection port of the adapter.</li>
	</ol>
	</li>
	<li>At co-ordinates AP: -3.08 mm ML: 0.5 mm, lower the 400 &#956;m diameter optic fiber into the VTA.
	<ol>
		<li>Before lowering the optic fiber, connect the ferrule to a fiber optic cable (with stainless steel or ceramic ferrule) linked to a&#160;fiber-coupled LED source. 
		NOTE: Make sure light intensity and focus are set as desired. The choice of the LED light source should correspond to the opsins being targeted. Here, a 470 nm (blue light) LED driver was used for hChR2 photoactivation (Figure 4a).</li>
		<li>To synchronize the light pulse with the neural recording, connect the LED driver and recording controller digital IN port to a transistor-transistor logic (TTL) pulser (Figure 4b). This can be achieved using a BNC splitter. 
		NOTE: The use of a TTL pulser provides the flexibility of digitally setting the desired pulse train frequency and duration. In the current protocol, 10 ms light pulses were triggered at 50 Hz18. For TTL control of the LED driver, switch the driver to &#8220;trigger&#8221;. In this mode, the light intensity can be regulated by turning the &#8220;knob&#8221;. Stimulation frequency can be adjusted to accommodate experimental variables&#160;and could range from 1 to 100 Hz. For neural circuit tracing, a stimulation frequency of &#62;20 Hz is recommended.</li>
		<li>Adjust the knob to determine the effective intensity that can generate a response without producing photoelectric artifacts. If necessary, reposition the optic fiber to eliminate artifacts19. 
		NOTE: The LED driver used for the current protocol produces light power of ~21.8 mW.</li>
	</ol>
	</li>
</ol>
5. Neural recording
<ol>
	<li>Use an acute neural probe with a 15&#8211;50 &#181;m thick shank, measuring at least 5 mm in length. 
	NOTE: Recording from deep brain structures will require neural probes with a longer shank. In the current study, a probe with 20 mm shank length was used.</li>
	<li>Connect the pre-amplifier head stage to the recording controller via a serial peripheral interface (SPI) cable. Check LEDs color lighting on the recording controller ports. Green and yellow LEDs indicate proper voltage on the connected amplifier board. Red LED indicates a working software-head stage control (Figure 5b).</li>
	<li>Using a micro-manipulator, position the electrode contact sites in the pyramidal cell layer of the CA1 (AP: -1.94mm, ML: +1.0mm, DV: +1.1 to 1.2mm). 
	NOTE: The optic fiber and electrode can be positioned in other desired brain regions for photostimulation and recording.</li>
</ol>
6. Amplifier and filter settings
<ol>
	<li>Connect the recording controller amplifier system to a computer through a USB 3.0 port. Connect the head stage to the amplifier using the SPI cable. Launch the controller software. If not properly connected, the system will display a &#8220;device not found&#8221; message.</li>
	<li>Click Run to view activity on the channels. Each waveform is plotted as the voltage (y-axis) versus time (x-axis). Depending on the region of interest, adjust the voltage and time scales to adjust the waveform display.</li>
	<li>If only a few channels are active, disable unused channels to reduce file size on the storage disk.</li>
	<li>Set the sampling rate and cut-off frequencies by editing bandwidth parameters on the acquisition software. For single-unit recording, set the upper and lower cutoff frequencies to 300 Hz and 5,000 Hz respectively. Change the amplifier sampling rate to 30 kS/s. 
	NOTE: Choosing a high sampling rate will produce a larger file size.</li>
	<li>Before recording, check the integrity of the electrode channels by measuring impedance at 1,000 Hz. This can be done by launching the function in the user interface (controller software). A working channel should have an impedance value that ranges from 0.1 to 5 M&#937;.</li>
	<li>If an audio output (speaker) is connected to ANALOG OUT port, adjust gain and silencer for optimum spike sound.</li>
	<li>To display the TTL pulse train marker in the spike recording, enable the display for the DIGITAL IN marker. To record the pulse train time stamp, enable the DIGITAL IN channel display before the main experiment. 
	NOTE: There is usually more than one &#8220;DIGITAL IN&#8221; channel (1 or 2). Ensure that the BNC cable from the TTL pulser is connected to the &#8220;DIGITAL IN&#8221; port that is selected for display in the recording controller software.</li>
	<li>For real-time viewing of the spike waveform, open the spike scope window, and set threshold using the mouse click.</li>
	<li>Inspect the noise level by monitoring the Root Mean Square (RMS in &#181;V). For clean spike recording, it is preferred that the neural spike threshold is at least 5x RMS.</li>
	<li>Once the setup is complete, select the file output format. 
	NOTE: Here, CA1 neural spike was recorded in .rhd format. Other file formats (.rhs and .dat) can also be selected to match the file formats permissible by the analysis software.</li>
</ol>
7. In vivo optogenetic recording and settings
<ol>
	<li>Open the pulse control software (TTL) which displays various adjustable pulse parameters. Select the appropriate COM port on the software. Set pulse parameters by adjusting the Train and Group settings. 
	NOTE: The pulse condition should be determined by considering experimental variables and design.</li>
	<li>On the amplifier controller software, click Run to detect neural spikes in the hippocampus or desired brain area. If necessary, adjust the electrode depth to detect viable and active neurons.</li>
	<li>Once extracellular voltage activity is detected, observe baseline spikes for 15 min and monitor the vitals of the animal. Also, test the light pulse to detect responsive neurons within the CA1 or desired anatomical target region. 
	NOTE: Check for photoelectric artifacts and eliminate by adjusting light intensity or optic fiber position.</li>
	<li>Record the baseline activity for ~10 min before triggering the light pulse at the desired frequency (Figure 5a,b). This will allow for comparison of firing or burst rates with and without illumination. 
	NOTE: In the current study, baseline activity was recorded for 10 min without illumination, followed by illumination at 50 Hz (Movie 1).</li>
</ol>
8. Analysis
<ol>
	<li>Convert&#160;the .rhd file into Nex5 output format.</li>
	<li>Process filename.Nex5 file in an offline sorter software to detect single unit raster trains and waveforms (Figure 5a&#8211;b).
	<ol>
		<li>Select the desired channel from the drop-down window. 
		NOTE: The continuously recorded spike data can be viewed on a separate window of the OFSS. The time scale can also be adjusted. If a tetrode is used, the OFSS can be set to sort the 4 channels as a tetrode.</li>
		<li>Set the voltage level for threshold crossing waveform extraction. 
		CAUTION: Use a minimum of 5x RMS for proper spike detection.</li>
		<li>Detect waveforms, then perform spike sorting using valley-seeking (automatic) or K-means (semi-automatic) method. Where necessary, merge units that occupy similar regions of the 3D-Principal Component Analysis (PCA) space.</li>
		<li>Export sorted .Nex5 files for further analysis (Figure 5c&#8211;d). 
		NOTE: Analysis outcomes for the interspike interval, firing rate, and burst rate can be exported into other analysis platforms.</li>
	</ol>
	</li>
</ol>
9. Fluorescence detection of AAV expression
<ol>
	<li>After the recording, euthanize the animal in an isoflurane chamber.</li>
	<li>Perform transcardial perfusion with 10 mM PBS (~10&#160;mL), followed by 4% phosphate-buffered paraformaldehyde (PB-PFA, ~10 mL).</li>
	<li>Remove the whole brain intact and fix it in 4% PB-PFA for 48 h.</li>
	<li>Transfer the fixed brain into 4% PB-PFA containing 30% sucrose for cryopreservation at 4 &#176;C. After 72 h, section the brain in a cryostat and mount slices on a gelatin-coated slide. 
	NOTE:&#160;In the current protocol, sections containing the closed part of the hippocampus (rostral) were selected to demonstrate AAV-labeled terminals in the DG, CA3, and CA1.</li>
</ol>

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+rewarding+and+novel+events+in+facilitating+memory+persistence+in+a+separate+spatial+memory+task.">The role of rewarding and novel events in facilitating memory persistence in a separate spatial memory task.</a> Learning &amp; Memory. 21 (2), 61-72 (2014).</li><li>Ntamati, N. R., Luscher, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=VTA+Projection+Neurons+Releasing+GABA+and+Glutamate+in+the+Dentate+Gyrus.">VTA Projection Neurons Releasing GABA and Glutamate in the Dentate Gyrus.</a> eNeuro. 3 (4), (2016).</li><li>Yoo, J. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ventral+tegmental+area+glutamate+neurons+co-release+GABA+and+promote+positive+reinforcement.">Ventral tegmental area glutamate neurons co-release GABA and promote positive reinforcement.</a> Nature Communications. 7, 13697 (2016).</li><li>Funahashi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Working+Memory+in+the+Prefrontal+Cortex.">Working Memory in the Prefrontal Cortex.</a> Brain Sciences. 7 (5), (2017).</li><li>Luo, A. H., Tahsili-Fahadan, P., Wise, R. A., Lupica, C. R., Aston-Jones, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Linking+context+with+reward:+a+functional+circuit+from+hippocampal+CA3+to+ventral+tegmental+area.">Linking context with reward: a functional circuit from hippocampal CA3 to ventral tegmental area.</a> Science. 333 (6040), 353-357 (2011).</li><li>McNamara, C. G., Dupret, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+sources+of+dopamine+for+the+hippocampus.">Two sources of dopamine for the hippocampus.</a> Trends in Neurosciences. 40 (7), 383-384 (2017).</li><li>McNamara, C. G., Tejero-Cantero, A., Trouche, S., Campo-Urriza, N., Dupret, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dopaminergic+neurons+promote+hippocampal+reactivation+and+spatial+memory+persistence.">Dopaminergic neurons promote hippocampal reactivation and spatial memory persistence.</a> Nature Neuroscience. 17 (12), 1658-1660 (2014).</li><li>Cardin, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+optogenetic+stimulation+and+recording+of+neurons+in+vivo+using+cell-type-specific+expression+of+Channelrhodopsin-2.">Targeted optogenetic stimulation and recording of neurons in vivo using cell-type-specific expression of Channelrhodopsin-2.</a> Nature Protocols. 5 (2), 247-254 (2010).</li><li>Mei, Y., Zhang, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+tools+and+approaches+for+optogenetics.">Molecular tools and approaches for optogenetics.</a> Biological Psychiatry. 71 (12), 1033-1038 (2012).</li><li>Zingg, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=AAV-Mediated+Anterograde+Transsynaptic+Tagging:+Mapping+Corticocollicular+Input-Defined+Neural+Pathways+for+Defense+Behaviors.">AAV-Mediated Anterograde Transsynaptic Tagging: Mapping Corticocollicular Input-Defined Neural Pathways for Defense Behaviors.</a> Neuron. 93 (1), 33-47 (2017).</li><li>Zhang, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+microbial+opsin+family+of+optogenetic+tools.">The microbial opsin family of optogenetic tools.</a> Cell. 147 (7), 1446-1457 (2011).</li><li>Aravanis, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optical+neural+interface:+in+vivo+control+of+rodent+motor+cortex+with+integrated+fiberoptic+and+optogenetic+technology.">An optical neural interface: in vivo control of rodent motor cortex with integrated fiberoptic and optogenetic technology.</a> Journal of Neural Engineering. 4 (3), 143-156 (2007).</li><li>Oron, D., Papagiakoumou, E., Anselmi, F., Emiliani, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+optogenetics.">Two-photon optogenetics.</a> Progress in Brain Research. 196, 119-143 (2012).</li></ol>

The authors have nothing to disclose.

In the past decade, the design of AAV constructs has advanced significantly. As such, more neuron-specific promoters have been incorporated into an array of AAV serotypes for improved transfection specificity14. By combining genes for fluorescence proteins, transporters, receptors, and ion channels, libraries of AAV now exists for imaging, neuromodulation, and synaptic activity detection. In commercially available AAV-constructs, a combination of a genetically encoded fluorophore and ion channels ...

In the past decade, an array of genetic tools was developed to increase the specificity of neuron-type modulation, and the mapping of complex neural networks1. Notably, neurotropic viruses with an inherent ability to infect and replicate in neuronal cells have been deployed to express or ablate specific proteins in neuron sub-types. When harboring fluorescence proteins or genetically encoded synaptic activity indicators, transfected AAV vectors label and delineate neural networks across brain regions2,3. The choice of a promoter in the AAV construct directs the expression of the vector in neuron types with some level of specificity (promoter-dependent expression). However, through Cre-lox recombination, AAV constructs are deployed with greater specificity for&#160;neuron labeling4,5,6,7. Of note, photoactivated microbial opsins and fluorescence proteins packaged in AAV vectors can be expressed in various neuron subtypes8, and are ideal for imaging, neuron-type circuit tracing, and photomodulation9,10.
AAVs constructs stereotactically injected into a brain region (or nucleus) drives the expression of the reporter protein in the soma, dendrite, and axons terminals. Neural expression of AAV harboring a reporter gene (eYFP) facilitates the labeling of neuron cell bodies and anatomical tracing of projections to and from other brain regions11,12,13,14. AAV-eYFP constructs carrying light-controlled opsin (e.g., hChR2), can be deployed as a tool for imaging6,15 and stimulation-based physiological tracing of neural projections to target brain areas in vivo16. Depending on the AAV serotype, the direction of neuron labeling may be anterograde or retrograde11,12. Previous studies have established that AAV5 travels anterogradely in neurons12. Thus, photostimulation&#160;of cell bodies expressing hChR2 produces presynaptic effects elsewhere in the brain (target)17.
Here, AAV (serotype 5) with a CaMKII&#945; promoter was used to express eYFP (reporter) and hChR2 (opsin) in VTA glutamate neurons and axonal projections. Results from this study demonstrate the layer-dependent distribution of VTA-glutamate presynaptic terminals in the CA1, CA3, and DG hippocampal regions. Also, photostimulation of VTA glutamate neurons increased CA1 multi-unit and single-unit firing rates in vivo when compared with baseline values. This protocol utilizes affordable tools and commercially available software that can increase the quality of data obtained from neural circuit tracing experiments.

<table>
	<tbody>
		<tr>
			<td>3% Hydrogen peroxide</td>
			<td>Fisher chemical</td>
			<td>H312</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV-CaMKII&#945;-ChR2-eGYP</td>
			<td>Addgene</td>
			<td>Plasmid #26969</td>
			<td></td>
		</tr>
		<tr>
			<td>BNC cable</td>
			<td>Amazon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BNC Splitter</td>
			<td>Amazon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ceramic Split Mating Sleeve for &#216;1.25mm Ferrules.</td>
			<td>Thorlabs</td>
			<td>ADAL1-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>Dremel</td>
			<td>LR 39098</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin coated slides</td>
			<td>Fisher scientific</td>
			<td>OBSLD01CS</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton&#39;s syringe (Neuros)</td>
			<td>WPI Inc.</td>
			<td>06H</td>
			<td></td>
		</tr>
		<tr>
			<td>Head stage adapter</td>
			<td>Neuronexus</td>
			<td>Adpt-Q4-OM32</td>
			<td></td>
		</tr>
		<tr>
			<td>High impedance silicon probe</td>
			<td>Neuronexus</td>
			<td>Q1x1-tet-5mm-121-CQ4</td>
			<td></td>
		</tr>
		<tr>
			<td>INTAN 512ch Recording Controller</td>
			<td>INTAN</td>
			<td>RHD2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Iodine solution</td>
			<td>Dynarex</td>
			<td>1425</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Piramal</td>
			<td>NDC 66794-017-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Spectrum</td>
			<td>K1068</td>
			<td></td>
		</tr>
		<tr>
			<td>LED Driver</td>
			<td>Thorlabs</td>
			<td>LEDD1B</td>
			<td></td>
		</tr>
		<tr>
			<td>LED light source (470 nm)-blue light</td>
			<td>Thorlabs</td>
			<td>M470F3</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Narishige</td>
			<td>M0-203</td>
			<td></td>
		</tr>
		<tr>
			<td>Optic fiber</td>
			<td>Thorlabs</td>
			<td>CFMLC14L05</td>
			<td></td>
		</tr>
		<tr>
			<td>Pan head philips screw (M0.6 X 2mm)</td>
			<td>Amazon</td>
			<td>M0.6 X 2mm</td>
			<td></td>
		</tr>
		<tr>
			<td>Pre-amplifier headstage (32 Channel)</td>
			<td>INTAN</td>
			<td>C3314</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic frame</td>
			<td>Kopf</td>
			<td>1530</td>
			<td></td>
		</tr>
		<tr>
			<td>TTL pulser</td>
			<td>Prizmatix</td>
			<td>4031</td>
			<td></td>
		</tr>
		<tr>
			<td>Urethane</td>
			<td>Sigma</td>
			<td>U2500</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Alfa Aesar</td>
			<td>J61430</td>
			<td></td>
		</tr>
		<tr>
			<td>Software</td>
			<td>Company</td>
			<td>Version</td>
			<td></td>
		</tr>
		<tr>
			<td>Graphpad Prism</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Intan Recording Controller</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Neuroexplorer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Plexon Offline Spike Sorter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ACSF Composition:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>oxygenated ACSF with 95% Oxygen/5%CO2 constantly being bubbled through the ACSF (ACSF; in mM 125 NaCl, 25 NaHCO3, 3 KCl, 1.25 NaH2PO4, 1 MgCl2, 2 CaCl2 and 25 Glucose).</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

combined in vivo anatomical and functional tracing of ventral tegmental area glutamate terminals in the hippocampus

Optogenetic modulation of neuron sub-populations in the brain has allowed researchers to dissect neural circuits in vivo and ex vivo. This provides a premise for determining the role of neuron types within a neural circuit, and their significance in information encoding relative to learning. Likewise, the method can be used to test the physiological significance of two or more connected brain regions in awake and anesthetized animals. The current study demonstrates how VTA glutamate neurons modulate the firing rate of putative pyramidal neurons in the CA1 (hippocampus) of anesthetized mice. This protocol employs adeno-associated virus (AAV)-dependent labeling of VTA glutamate neurons for the tracing of VTA presynaptic glutamate terminals in the layers of the hippocampus. Expression of light-controlled opsin (channelrhodopsin; hChR2) and fluorescence protein (eYFP) harbored by the AAV vector permitted anterograde tracing of VTA glutamate terminals, and photostimulation of VTA glutamate neuron cell bodies (in the VTA). High-impedance acute silicon electrodes were positioned in the CA1 to detect multi-unit and single-unit responses to VTA photostimulation in vivo. The results of this study demonstrate the&#160;layer-dependent distribution of presynaptic VTA glutamate terminals in the hippocampus (CA1, CA3, and DG). Also, the photostimulation of VTA glutamate neurons increased the firing and burst rate of putative CA1 pyramidal units in vivo.

Anterograde tracing
AAV expression was verified by immunofluorescence imaging of reporter protein (eYFP) in the VTA of C57BL/6 mice 21 days post-injection (Figure 2). Successful anterograde labeling of presynaptic VTA glutamate projections in the hippocampus was also verified by eYFP detection in the layers of the DG, CA3, and CA1 (Figure 6a&#8211;d; Movie 2 and 3).
<strong...

Watch this Scientific Journal Video about Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus at JoVE.com

Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus

The current protocol demonstrates a simple method for tracing of ventral tegmental area (VTA) glutamate projections to the hippocampus. Photostimulation of VTA glutamate neurons was combined with CA1 recording to demonstrate how VTA glutamate terminals modulate CA1 putative pyramidal firing rate in vivo.

Optogenetic modulation of neuron sub-populations in the brain has allowed researchers to dissect neural circuits in vivo and ex vivo. This provides a ...

combined-vivo-anatomical-functional-tracing-ventral-tegmental-area

Research

JoVE Journal

Neuroscience

3.7K Views.  Louisiana State University School of Veterinary Medicine. The current protocol demonstrates a simple method for tracing of ventral tegmental area (VTA) glutamate projections to the hippocampus. Photostimulation of VTA glutamate neurons was combined with CA1 recording to demonstrate how VTA glutamate terminals modulate CA1 putative pyramidal firing rate in vivo.

Video: Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Comprehensive Profiling of Dopamine Regulation in Substantia Nigra and Ventral Tegmental Area

Dopamine is a vigorously studied neurotransmitter in the CNS. Indeed, its involvement in locomotor activity and reward-related behaviour has fostered five decades of inquiry into the molecular deficiencies associated with dopamine regulation. The majority of these inquiries of dopamine regulation in the brain focus upon the molecular basis for its regulation in the terminal field regions of the nigrostriatal and mesoaccumbens pathways; striatum and nucleus accumbens. Furthermore, such studies have concentrated on analysis of dopamine tissue content with normalization to only wet tissue weight. Investigation of the proteins that regulate dopamine, such as tyrosine hydroxylase (TH) protein, TH phosphorylation, dopamine transporter (DAT), and vesicular monoamine transporter 2 (VMAT2) protein often do not include analysis of dopamine tissue content in the same sample. The ability to analyze both dopamine tissue content and its regulating proteins (including post-translational modifications) not only gives inherent power to interpreting the relationship of dopamine with the protein level and function of TH, DAT, or VMAT2, but also extends sample economy. This translates into less cost, and yet produces insights into the molecular regulation of dopamine in virtually any paradigm of the investigators' choice.
We focus the analyses in the midbrain. Although the SN and VTA are typically neglected in most studies of dopamine regulation, these nuclei are easily dissected with practice. A comprehensive readout of dopamine tissue content and TH, DAT, or VMAT2 can be conducted. There is burgeoning literature on the impact of dopamine function in the SN and VTA on behavior, and the impingements of exogenous substances or disease processes therein 1-5. Furthermore, compounds such as growth factors have a profound effect on dopamine and dopamine-regulating proteins, to a comparatively greater extent in the SN or VTA 6-8. Therefore, this methodology is presented for reference to laboratories that want to extend their inquiries on how specific treatments modulate behaviour and dopamine regulation. Here, a multi-step method is presented for the analyses of dopamine tissue content, the protein levels of TH, DAT, or VMAT2, and TH phosphorylation from the substantia nigra and VTA from rodent midbrain. The analysis of TH phosphorylation can yield significant insights into not only how TH activity is regulated, but also the signaling cascades affected in the somatodendritic nuclei in a given paradigm.
We will illustrate the dissection technique to segregate these two nuclei and the sample processing of dissected tissue that produces a profile revealing molecular mechanisms of dopamine regulation in vivo, specific for each nuclei (Figure 1).

Dopamine is distinctly regulated in the midbrain nuclei, which contain the cell bodies and dendrites of the dopamine neurons. Here we describe a dissection and sample-handling approach to maximize results, and thus conclusions and insights, on dopamine regulation in the midbrain nuclei of the substantia nigra (SN) and ventral tegmental area (VTA) in rodents.

Dopamine is a vigorously studied neurotransmitter in the CNS. Indeed, its involvement in locomotor activity and reward-related behaviour has fostered five ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Laser Capture Microdissection - A Demonstration of the Isolation of Individual Dopamine Neurons and the Entire Ventral Tegmental Area

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue sections on a microscope slide. When combined with immunohistochemistry, LCM can be used to isolate individual cells types based on a specific protein marker. Here, the LCM technique is described for collecting a specific population of dopamine neurons directly labeled with&#160;tyrosine hydroxylase immunohistochemistry and for isolation of the dopamine neuron containing region of the ventral tegmental area using indirect tyrosine hydroxylase immunohistochemistry on a section adjacent to those used for LCM. An infrared (IR) capture laser is used to both dissect individual neurons as well as the ventral tegmental area off glass slides and onto an LCM cap for analysis. Complete dehydration of the tissue with 100% ethanol and xylene is critical. The combination of the IR capture laser and the ultraviolet (UV) cutting laser is used to isolate individual dopamine neurons or the ventral tegmental area when using PEN membrane slides. A PEN membrane slide has significant advantages over a glass slide as it offers better consistency in capturing and collecting cells, is faster collecting large pieces of tissue, is less reliant on dehydration and results in complete removal of the tissue from the slide. Although removal of large areas of tissue from a glass slide is feasible, it is considerably more time consuming and frequently leaves some residual tissue behind. Data shown here demonstrate that RNA of sufficient quantity and quality can be obtained using these procedures for quantitative PCR measurements. Although RNA and DNA are the most commonly isolated molecules from tissue and cells collected with LCM, isolation and measurement of microRNA, protein and epigenetic changes in DNA can also benefit from the enhanced anatomical and cellular resolution obtained using LCM.

The isolation of individual dopamine neurons or the ventral tegmental area with direct or indirect immunohistochemistry is demonstrated using laser capture microdissection. Parameters for isolation of tissue from a glass slide using an infrared laser and from membrane slides using the combination of an infrared and ultraviolet laser are discussed.

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue ...

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.

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 ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new method of Ca2+-imaging using the bioluminescent reporter GFP-aequorin (GA) has been developed. This new technique relies on the fusion of the GFP and aequorin genes, producing a molecule capable of binding calcium and&#160;&#8212; with the addition of its cofactor coelenterazine &#8212; emitting bright light that can be monitored through a photon collector. Transgenic lines carrying the GFP-aequorin gene have been generated for both mice and Drosophila. In Drosophila, the GFP-aequorin gene has been placed under the control of the GAL4/UAS binary expression system allowing for targeted expression and imaging within the brain. This method has subsequently been shown to be capable of detecting both inward Ca2+-transients and Ca2+-released from inner stores. Most importantly it allows for a greater duration in continuous recording, imaging at greater depths within the brain, and recording at high temporal resolutions (up to 8.3 msec). Here we present the basic method for using bioluminescent imaging to record and analyze Ca2+-activity within the mushroom bodies, a structure central to learning and memory in the fly brain.

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Here we present a novel Ca2+-imaging approach using a bioluminescent reporter. This approach uses a fused construct GFP-aequorin which binds to Ca2+ and emits light, eliminating the need for light excitation. Significantly this method permits long continuous imaging, access to deep brain structures and high temporal resolution.

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate Receptors in the Mouse Amygdala

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to study the molecular composition of membranes produced a significant decline in its use. Recently, there has been a major revival in freeze-fracture electron microscopy thanks to the development of effective ways to reveal integral membrane proteins by immunogold labeling. One of these methods is known as detergent-solubilized Freeze-fracture Replica Immunolabeling (FRIL).
The combination of the FRIL technique with optogenetics allows a correlated analysis of the structural and functional properties of central synapses. Using this approach it is possible to identify and characterize both pre- and postsynaptic neurons by their respective expression of a tagged channelrhodopsin and specific molecular markers. The distinctive appearance of the postsynaptic membrane specialization of glutamatergic synapses further allows, upon labeling of ionotropic glutamate receptors, to quantify and analyze the intrasynaptic distribution of these receptors. Here, we give a step-by-step description of the procedures required to prepare paired replicas and how to immunolabel them. We will also discuss the caveats and limitations of the FRIL technique, in particular those associated with potential sampling biases. The high reproducibility and versatility of the FRIL technique, when combined with optogenetics, offers a very powerful approach for the characterization of different aspects of synaptic transmission at identified neuronal microcircuits in the brain.
Here, we provide an example how this approach was used to gain insights into structure-function relationships of excitatory synapses at neurons of the intercalated cell masses of the mouse amygdala. In particular, we have investigated the expression of ionotropic glutamate receptors at identified inputs originated from the thalamic posterior intralaminar and medial geniculate nuclei. These synapses were shown to relay sensory information relevant for fear learning and to undergo plastic changes upon fear conditioning.

This article illustrates how the expression of neurotransmitter receptors can be quantified and the pattern analyzed at synapses with identified pre and postsynaptic elements using a combination of viral transduction of optogenetic tools and the freeze-fracture replica immunolabeling technique.

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to ...