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&amp;alpha;-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 &amp;Oslash;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&#039;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>&lt;strong&gt;Software&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Version&lt;/strong&gt;</td><td></td></tr><tr><td>Graphpad Prism</td><td>GraphPad Software</td><td>version 8</td><td></td></tr><tr><td>Intan Recording Controller</td><td>Intantech</td><td>version 2.07</td><td></td></tr><tr><td>Neuroexplorer</td><td>Nex Technologies</td><td>version 5.215</td><td></td></tr><tr><td>Plexon Offline Spike Sorter</td><td>Plexon Inc</td><td>version 4.5</td><td></td></tr><tr><td>&lt;strong&gt;ACSF Composition:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>oxygenated ACSF with 95% Oxygen/5%CO&lt;sub&gt;2&lt;/sub&gt; constantly being bubbled through the ACSF (ACSF; in mM 125 NaCl, 25 NaHCO&lt;sub&gt;3&lt;/sub&gt;, 3 KCl, 1.25 NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 1 MgCl&lt;sub&gt;2&lt;/sub&gt;, 2 CaCl&lt;sub&gt;2&lt;/sub&gt; 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 ...

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Research

JoVE Journal

Neuroscience

3.8K 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

Longitudinal Two-Photon Imaging of Dorsal Hippocampal CA1 in Live Mice

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from subcellular to network levels and at temporal scales from milliseconds to weeks. In addition, two-photon imaging can be combined with a variety of behavioral tasks to explore the causal relationships between brain function and behavior. However, in mammals, limited penetration and scattering of light have limited two-photon intravital imaging mostly to superficial brain regions, thus precluding longitudinal investigation of deep-brain areas such as the hippocampus. The hippocampus is involved in spatial navigation and episodic memory and is a long-standing model used to study cellular as well as cognitive processes important for learning and recall, both in health and disease. Here, a preparation that enables chronic optical access to the dorsal hippocampus in living mice is detailed. This preparation can be combined with two-photon optical imaging at cellular and subcellular resolution in head fixed, anesthetized live mice over several weeks. These techniques enable repeated imaging of neuronal structure or activity-evoked plasticity in tens to hundreds of neurons in the dorsal hippocampal CA1. Furthermore, this chronic preparation can be used in combination with other techniques such as micro-endoscopy, head-mounted wide field microscopy or three-photon microscopy, thus greatly expanding the toolbox to study cellular and network processes involved in learning and memory.

This method describes a chronic preparation that allows optical access to the hippocampus of living mice. This preparation can be used to perform longitudinal optical imaging of neuronal structural plasticity and activity-evoked cellular plasticity over a period of several weeks.

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from ...

Behavior

Measurement of Glutamate Uptake using Radiolabeled L-[3H]-Glutamate in Acute Transverse Slices Obtained from Rodent Resected Hippocampus

Glutamate removal from the extracellular space by high-affinity Na+-dependent transporters is essential to ensure that the brain&#39;s intrinsic connectivity mechanisms work properly and homeostasis is maintained. The hippocampus is a unique brain structure that manages higher cognitive functions, and is the subject of several studies regarding neurologic diseases. The investigation of physiological and pathological mechanisms in rodent models can benefit from acute hippocampal slice (AHS) preparations. AHS has the advantage of providing reliable information on cell function since the cytoarchitecture and synaptic circuits are preserved. Although AHS preparations are commonly used in neurochemistry laboratories, it is possible to find some methodological differences in the literature. Considering that distinctive slice preparation protocols might change the hippocampal regions analyzed, this current protocol proposes a standard technique for obtaining transverse AHS from resected hippocampus. This simple-to-perform protocol may be used in mice and rats&#39; experimental models and allow several ex vivo approaches investigating neurochemical dynamics (in dorsal, intermediate and ventral hippocampus) in different backgrounds (e.g., transgenic manipulations) or after in vivo manipulations (e.g., pharmacological treatments or suitable rodent models to study clinical disorders). After dissecting the hippocampus from the rodent brain, transverse slices along the septo-temporal axis (300 &#181;m thick) were obtained. These AHS contain distinct parts of the hippocampus and were subjected to an individual neurochemical investigation (as an example: neurotransmitter transporters using their respective substrates). As the hippocampus presents a high density of excitatory synapses, and glutamate is the most important neurotransmitter in the brain, the glutamatergic system is an interesting target for in vivo observed phenomena. Thus, the current protocol provides detailed steps to explore glutamate uptake in ex vivo AHS using L-[3H]-Glutamate. Using this protocol to investigate hippocampal function may help to better understand the influence of glutamate metabolism on mechanisms of neuroprotection or neurotoxicity.

Measurement of Glutamate Uptake using Radiolabeled L-[3H]-Glutamate in Acute Transverse Slices Obtained from Rodent Resected Hippocampus

This article describes a reliable and simple way to obtain ex vivo acute hippocampal transverse slices from mice and rats using a tissue chopper. Slices obtained from resected hippocampi can be submitted for functional glutamate uptake analysis to investigate glutamatergic system homeostasis.

Glutamate removal from the extracellular space by high-affinity Na+-dependent transporters is essential to ensure that the brain&#39;s intrinsic ...

Biochemistry

High-resolution In Vivo Manual Segmentation Protocol for Human Hippocampal Subfields Using 3T Magnetic Resonance Imaging

The human hippocampus has been broadly studied in the context of memory and normal brain function and its role in different neuropsychiatric disorders has been heavily studied. While many imaging studies treat the hippocampus as a single unitary neuroanatomical structure, it is, in fact, composed of several subfields that have a complex three-dimensional geometry. As such, it is known that these subfields perform specialized functions and are differentially affected through the course of different disease states. Magnetic resonance (MR) imaging can be used as a powerful tool to interrogate the morphology of the hippocampus and its subfields. Many groups use advanced imaging software and hardware (&#62;3T) to image the subfields; however this type of technology may not be readily available in most research and clinical imaging centers. To address this need, this manuscript provides a detailed step-by-step protocol for segmenting the full anterior-posterior length of the hippocampus and its subfields: cornu ammonis (CA) 1, CA2/CA3, CA4/dentate gyrus (DG), strata radiatum/lacunosum/moleculare (SR/SL/SM), and subiculum. This protocol has been applied to five subjects (3F, 2M; age 29-57, avg. 37). Protocol reliability is assessed by resegmenting either the right or left hippocampus of each subject and computing the overlap using the Dice&#39;s kappa metric. Mean Dice&#39;s kappa (range) across the five subjects are: whole hippocampus, 0.91 (0.90-0.92); CA1, 0.78 (0.77-0.79); CA2/CA3, 0.64 (0.56-0.73); CA4/dentate gyrus, 0.83 (0.81-0.85); strata radiatum/lacunosum/moleculare, 0.71 (0.68-0.73); and subiculum 0.75 (0.72-0.78). The segmentation protocol presented here provides other laboratories with a reliable method to study the hippocampus and hippocampal subfields in vivo using commonly available MR tools.

High-resolution In Vivo Manual Segmentation Protocol for Human Hippocampal Subfields Using 3T Magnetic Resonance Imaging

The goal of this manuscript is to study the hippocampus and hippocampal subfields using MRI. The manuscript describes a protocol for segmenting the hippocampus and five hippocampal substructures: cornu ammonis (CA) 1, CA2/CA3, CA4/dentate gyrus, strata radiatum/lacunosum/moleculare, and subiculum.

The human hippocampus has been broadly studied in the context of memory and normal brain function and its role in different neuropsychiatric disorders has ...

Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, it is widely used to study activity-related calcium signals in small subcellular compartments such as dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces postsynaptic sodium signals, the properties of which are only marginally understood. Here, we describe a method for combined whole-cell patch-clamp and multi-photon sodium imaging in cellular micro domains of central neurons. Furthermore, we introduce a modified procedure for ultra-violet (UV)-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue. To this end, whole-cell recordings were performed on Cornu Ammonis subdivision 1 (CA1) pyramidal neurons in acute tissue slices of the mouse hippocampus. Neurons were filled with the sodium-sensitive fluorescent dye SBFI through the patch-pipette, and multi-photon excitation of SBFI enabled the visualization of dendrites and adjacent spines. To establish UV-induced focal uncaging, several parameters including light intensity, volume affected by the UV uncaging beam, positioning of the beam as well as concentration of the caged compound were tested and optimized. Our results show that local perfusion with caged glutamate (MNI-Glutamate) and its focal UV-uncaging result in inward currents and sodium transients in dendrites and spines. Time course and amplitude of both inward currents and sodium signals correlate with the duration of the uncaging pulse. Furthermore, our results show that intracellular sodium signals are blocked in the presence of blockers for ionotropic glutamate receptors, demonstrating that they are mediated by sodium influx though this pathway. In summary, our method provides a reliable tool for the investigation of intracellular sodium signals induced by focal receptor activation in intact brain tissue.

We describe the combination of focal UV-induced photo-activation of neuro-active compounds with whole-cell patch-clamp and multi-photon imaging of intracellular sodium transients in dendrites and spines of hippocampal neurons in acute tissue slices of the mouse brain.

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high ...

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 ...

Improving the Application of High Molecular Weight Biotinylated Dextran Amine for Thalamocortical Projection Tracing in the Rat

High molecular weight biotinylated dextran amine (BDA) has been used as a highly sensitive neuroanatomical tracer for many decades. Since the quality of its labeling was affected by various factors, here, we provide a refined protocol for the application of high molecular weight BDA for studying optimal neural labeling in the central nervous system. After stereotactic injection of BDA into the ventral posteromedial nucleus (VPM) of the thalamus in the rat through a delicate glass pipette, BDA was stained with fluorescent streptavidin-Alexa (AF) 594 and counterstained with fluorescent Nissl stain AF500/525. On the background of green Nissl staining, the red BDA labeling, including neuronal cell bodies and axonal terminals, was more distinctly demonstrated in the somatosensory cortex. Furthermore, double fluorescent staining for BDA and the calcium-binding protein parvalbumin (PV) was carried out to observe the correlation of BDA labeling and PV-positive interneurons in the cortical target, providing the opportunity to study the local neural circuits and their chemical characteristics. Thus, this refined method is not only suitable for visualizing high quality neural labeling with the high molecular weight BDA through reciprocal neural pathways between the thalamus and cerebral cortex, but also will permit the simultaneous demonstration of other neural markers with fluorescent histochemistry or immunochemistry.

Here, we present a refined protocol to effectively reveal biotinylated dextran amine (BDA) labeling with a fluorescent staining method through a reciprocal neural pathway. It is suitable for analyzing the fine structure of BDA labeling and distinguishing it from other neural elements under a confocal laser scanning microscope.

High molecular weight biotinylated dextran amine (BDA) has been used as a highly sensitive neuroanatomical tracer for many decades. Since the quality of ...

In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional investigation of long-range connections requires targeted recordings of single neurons combined with the specific stimulation of identified distant inputs. This is often difficult to achieve with conventional and electrical stimulation techniques, because axons from converging upstream brain areas may intermingle in the target region. The stereotaxic targeting of a specific brain region for virus-mediated expression of light-sensitive ion channels allows selective stimulation of axons originating from that region with light. Intracerebral stereotaxic injections can be used in well-delimited structures, such as the anterior thalamic nuclei, in addition to other subcortical or cortical areas throughout the brain.
Described here is a set of techniques for precise stereotaxic injection of viral vectors expressing channelrhodopsin in the mouse brain, followed by photostimulation of axon terminals in the brain slice preparation. These protocols are simple and widely applicable. In combination with whole-cell patch clamp recording from a postsynaptically connected neuron, photostimulation of axons allows the detection of functional synaptic connections, pharmacological characterization, and evaluation of their strength. In addition, biocytin filling of the recorded neuron can be used for post-hoc morphological identification of the postsynaptic neuron.

This protocol describes a set of methods to identify the cell-type specific functional connectivity of long-range inputs from distant brain regions using optogenetic stimulations in ex vivo brain slices.

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional ...

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

Microglia, the only immune cells resident in the brain, actively participate in neural circuit maintenance by modifying synapses and neuronal excitability. Recent studies have revealed the differential gene expression and functional heterogeneity of microglia in different brain regions. The unique functions of the hippocampal neural network in learning and memory may be associated with the active roles of microglia in synapse remodeling. However, inflammatory responses induced by surgical procedures have been problematic in the two-photon microscopic analysis of hippocampal microglia. Here, a method is presented that enables the chronic observation of microglia in all layers of the hippocampal CA1 through an imaging window. This method allows the analysis of morphological changes in microglial processes for more than 1 month. Long-term and high-resolution imaging of the resting microglia requires minimally invasive surgical procedures, appropriate objective lens selection, and optimized imaging techniques. The transient inflammatory response of hippocampal microglia may prevent imaging immediately after surgery, but the microglia restore their quiescent morphology within a few weeks. Furthermore, imaging neurons simultaneously with microglia allows us to analyze the interactions of multiple cell types in the hippocampus. This technique may provide essential information about microglial function in the hippocampus.

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

This paper describes a method for chronic in vivo observation of the resting microglia in the mouse hippocampal CA1 using precisely controlled surgery and two-photon microscopy.

Microglia, the only immune cells resident in the brain, actively participate in neural circuit maintenance by modifying synapses and neuronal ...

Real-Time Imaging for Neuron Activity in the Dentate Gyrus Cells of Mice 

In Vivo Calcium Imaging of Granule Cells in the Dentate Gyrus of Hippocampus in Mice

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal activity in awake animals during behavior tasks. Since the hippocampus is closely associated with episodic and spatial memory, it has become an essential brain region in&#160;this field&#39;s research. In recent research, engram cells and place cells were studied by recording the neural activities in the hippocampal CA1 region using the miniature microscope in mice while performing behavioral tasks including open-field and linear track. Although the dentate gyrus is another important region in the hippocampus, it has rarely been studied with in vivo imaging due to its greater depth and difficulty for imaging. In this protocol, we present in detail a calcium imaging process, including how to inject the virus, implant a GRIN (Gradient-index) lens, and attach a base plate for imaging the dentate gyrus of the hippocampus. We further describe how to preprocess the calcium imaging data using MATLAB. Additionally, studies of other deep brain regions that require imaging may benefit from this method.

Author Spotlight: Investigating Neural Activity of Dentate Gyrus Granule Cells with Miniature Microscope

The dentate gyrus of the hippocampus carries out essential and distinct functions in learning and memory. This protocol describes a set of robust and efficient procedures for in vivo calcium imaging of granule cells in the dentate gyrus in freely moving mice.

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal ...

Co-culturing of Ventral Hippocampal and Nucleus Accumbens

Source: Zhong, C., et al. <a href="https://app.jove.com/v/52730">Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons.</a> J. Vis. Exp. (2015)
This video demonstrates the process of preparing ventral hippocampal and nucleus accumbens tissues from mouse brains for co-culture. Neurons extend projections to form synaptic connections between the ventral hippocampal and nucleus accumbens tissues. This method allows studying neuronal interactions between these two brain regions, offering insights into cognition, behavior, and neurological disorders.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Combined In Vivo Anatomical and Functional Tracing of Ventral Tegmental Area Glutamate Terminals in the Hippocampus

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Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

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

Improving the Application of High Molecular Weight Biotinylated Dextran Amine for Thalamocortical Projection Tracing in the Rat

In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

In Vivo Calcium Imaging of Granule Cells in the Dentate Gyrus of Hippocampus in Mice

Co-culturing of Ventral Hippocampal and Nucleus Accumbens