This work was supported by a Deutsche Forschungsgemeinschaft Research Fellowship (GO 3060/1-1, project number 401540516, to DG) and National Institutes of Health grant R56 DC016880 (MTR).

Obtain approval from the local Institutional Animal Care and Use Committee (IACUC) and adhere to NIH guidelines for the care and use of laboratory animals. All procedures in this protocol were approved by the University of Michigan IACUC and were in accordance with NIH guidelines for the care and use of laboratory animals.
1. Surgery Preparations
<ol>
	<li>Perform surgeries in aseptic conditions. Autoclave/sterilize all surgery tools and materials before surgery. Wear surgery gown and mask for the surgery.</li>
	<li>Sanitize the surgery area (spray and wipe down with 70% ethanol), and place sterile towel drapes to cover the surgery area.</li>
	<li>Prepare the recovery cage. Remove cage bedding to limit risk of asphyxiation. Put a heating pad under cage. Provide a food and water source.</li>
	<li>Pull a glass capillary for the nanoinjector on a pipette puller. The intense heat of the heating filament during the pulling process will sterilize the glass capillary. Cut or break off the tip to obtain an opening approximately 5 &#181;m in diameter.</li>
	<li>Bevel the capillary tip to an approximately 30&#176; angle to improve tissue penetration and reduce clogging. Backfill the capillary with mineral oil and insert into a nanoliter injector.</li>
	<li>Obtain an aliquot of the desired channelrhodopsin-encoding rAAV and dilute to the desired titer using sterile PBS. 
	NOTE: We have found that serotype 1 rAAVs work well for transfection of auditory brainstem nuclei. Specifically, rAAV1.Syn.Chronos-GFP.WPRE.bGH (blue light-activated channelrhodopsin) and rAAV1.Syn.ChrimsonR-tdTomato.WPRE.bGH (red-shifted channelrhodopsin), which are available from publically accessible repositories and vector cores, consistently yield the high expression levels and good long-range axonal trafficking of channelrhodopsins needed for CRACM experiments.</li>
	<li>Follow injector instructions to front fill capillary with 1-3 &#181;L of rAAV in sterile PBS.</li>
</ol>
2. Surgery
<ol>
	<li>Put the animal into an induction chamber and induce anesthesia with 3% isoflurane in oxygen delivered via a calibrated isoflurane vaporizer. Observe the mouse until breathing becomes deep and slow and a toe pinch reflex is absent, about 3-5 min.</li>
	<li>Transfer the animal to a stereotaxic frame. Secure the animal&#39;s head by putting its mouth on a palate bar with a gas anesthesia mask and by positioning non-perforating ear bars in both ear canals.</li>
	<li>Insert a rectal temperature probe and switch on the homeostatic temperature controller.</li>
	<li>Apply ophthalmic ointment to prevent eyes from drying out.</li>
	<li>Administer preemptive analgesic (e.g., subcutaneous injection of 5 mg/kg carprofen).</li>
	<li>Adjust isoflurane to 1-2.5%, according to the depth of the anesthetized state. Monitor temperature, breathing and color of mucous membranes at least every 15 minutes during the procedure.</li>
	<li>Shave scalp with electric clippers. Aseptically prepare scalp with three alternating swabs of povidone-iodine and 70% ethanol.</li>
	<li>Make an incision in the scalp along the midline starting between the ears and continuing rostral to the eyes, exposing the lambda and bregma sutures. Push skin to the side and remove periosteum from exposed bone if necessary.</li>
	<li>Mark the lambda suture with a sterile surgical marker, position the tip of the nanoinjector so that it is just touching lambda, and zero the micromanipulator coordinates. Use the nanoinjector tip and micromanipulator to measure the difference in elevation between the lambda and bregma sutures. Adjust palate bar height to bring lambda and bregma to within &#177; 100 &#181;m height difference.</li>
	<li>Map the injection site using the nanoinjector tip and micromanipulator coordinate system and mark the site with a sterile surgical marker. To inject the IC or DCN of P21-P30 mice, use coordinates relative to the lambda suture, as shown in Table 1. Note that the Z depth in our coordinates is measured from the surface of the skull at lambda.</li>
	<li>Use a micromotor drill with a sterile 0.5 mm drill burr to perform a craniotomy over the injection site.</li>
	<li>To ensure broad transfection of neurons in the target nucleus, make injections at various depths into the tissue (Table 1, Z coordinates), and, in the case of larger brain regions like the IC, make injections over the course of two or more penetrations at different X and Y coordinates (Table 1, Right IC penetration 1 and Right IC penetration 2).</li>
	<li>Perform injections. For IC injections, deposit 20 nL of virus in intervals of 250 &#181;m along the Z axis (injection depth) between 2,250 &#181;m and 1750 &#181;m depth. For DCN injections, deposit 20 nL of virus at a depth of 4,750 &#181;m and 4,550 &#181;m, respectively.</li>
	<li>After injection at each Z coordinate, wait 2-3 min before moving the injector to the next Z coordinate. This will allow time for the virus to diffuse away from the injection site, reducing the probability that virus will be sucked up the injection tract when the nanoinjector is repositioned.</li>
	<li>After last injection in a penetration, wait 3-5 min before retracting nanoinjector from brain.</li>
	<li>When the nanoinjector is removed from the brain between penetrations and between animals, eject a small volume of virus from the tip to check that the tip has not clogged.</li>
	<li>After injections, use sterile PBS to wet the cut edges of the scalp and then gently move the skin back towards the midline. Close the wound with simple interrupted sutures using 6-0 (0.7 metric) nylon sutures.</li>
	<li>Apply 0.5-1 mL of 2% lidocaine gel to the wound.</li>
	<li>Remove ear bars and temperature probe, turn off isoflurane, remove the mouse from the palate bar and transfer it to the recovery cage.</li>
	<li>Monitor recovery closely. Once the animal is fully awake, moving around, and showing no signs of pain or distress, transfer it back into its cage and return the cage to the vivarium.</li>
	<li>If surgeries will be performed on multiple animals in one day, use a hot bead sterilizer to sanitize surgery tools and drill burr before next surgery.</li>
</ol>
3. Surgical Follow Up
<ol>
	<li>Check animals daily for wound closure, infection, or signs of pain or distress over the next 10 days, adhering to the institution&#39;s animal care guidelines.</li>
	<li>Wait 3-4 weeks before using animals in experiments to allow optimal expression of the channelrhodopsins.</li>
</ol>
4. Brain Slice Preparation and Confirmation of Injection Target
<ol>
	<li>For CRACM, use acutely prepared brain slices from transfected animals in standard in vitro electrophysiology experiments, described here only briefly (see Goyer et al. 2019 for a more detailed description27).</li>
	<li>Prepare artificial cerebrospinal fluid (ACSF) containing (in mM): 125 NaCl, 12.5 D-glucose, 25 NaHCO3, 3 KCl, 1.25 NaH2PO4, 1.5 CaCl2, 1 MgSO4. Bubble ACSF to a pH of 7.4 with 5% CO2 in 95% O2.</li>
	<li>Perform all following steps, including in vitro electrophysiology, in near-darkness or red light to limit activation of channelrhodopsins.</li>
	<li>Deeply anesthetize mouse with isoflurane and decapitate it quickly. Dissect the brain quickly in ~34 &#176;C ACSF.</li>
	<li>Cut coronal slices (200-250 &#181;m) containing the IC in ~34 &#176;C ACSF with a vibrating microtome and incubate the slices at 34 &#176;C for 30 min in a holding chamber filled with ACSF bubbled with 5% CO2 in 95% O2. After incubation, store slices at room temperature until used for recordings.</li>
	<li>If the injection target was not the IC, cut additional coronal slices of the injected brain region and check the transfection of the target nucleus under a fluorescence microscope. If there is no transfection in the target nucleus or additional transfection in different brain regions, do not continue with experiment.</li>
</ol>
5. In Vitro Recording and CRACM Experiment
NOTE: To provide optical stimulation of Chronos and ChrimsonR, we use LEDs coupled to the epifluorescence port of the microscope. However, lasers can be used instead of LEDs. If using lasers, obtain prior approval from institutional safety officials and follow appropriate guidelines for safe laser use.
<ol>
	<li>Pull electrodes from borosilicate glass to a resistance of 3.5-4.5 M&#937;. The electrode internal solution should contain (in mM): 115 K-gluconate, 7.73 KCl, 0.5 EGTA, 10 HEPES, 10 Na2 phosphocreatine, 4 MgATP, 0.3 NaGTP, supplemented with 0.1% biocytin (w/v), pH adjusted to 7.3 with KOH and osmolality to 290 mOsm/kg with sucrose.</li>
	<li>To make recordings, use standard patch clamp methods. Place the slice in a recording chamber under a fixed stage upright microscope and continuously perfuse with ACSF at ~2 mL/min. Conduct recordings near physiological temperature (~34-36 &#176;C).</li>
	<li>Patch neurons under visual control using a suitable patch clamp amplifier. Correct for series resistance, pipette capacitance and liquid junction potential.</li>
	<li>During whole cell recordings, activate Chronos by delivering brief pulses (1-5 ms) of 470 nm light or ChrimsonR by brief pulses of 580 nm light through commercially available LEDs. Determine threshold of opsin activation and use a minimal stimulation protocol to elicit postsynaptic potentials. In general, use the shortest stimulus duration that elicits a PSP, and set the optical power to 120% of the threshold power required to elicit PSPs.</li>
	<li>To confirm that the recorded changes in membrane potential are indeed synaptic inputs to the neuron, standard antagonists for excitatory/inhibitory postsynaptic receptors can be washed in during the experiment. To investigate different receptor contributions to a PSP (e.g. NMDA vs AMPA receptors), suitable receptor antagonists can be washed in. For each receptor antagonist, drug effects should reverse after washout.</li>
	<li>Use the latency, jitter, and reliability of PSPs to confirm that light-activated synaptic inputs originate from direct, optical activation of synapses on the recorded neuron, as opposed to activation of channelrhodopsin-expressing synapses on an intervening neuron that synapses on the recorded neuron. In general, low latency (&#60;2 ms), low jitter (&#60;1 ms standard deviation in latency), and high reliability (&#62;50%) indicate a direct synaptic connection from the channelrhodopsin expressing presynaptic neuron to the recorded neuron.</li>
</ol>

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The authors have nothing to disclose.

We have found that CRACM is a powerful technique for identifying and characterizing long range synaptic inputs to neurons in the mouse IC. Following the protocol detailed here, we achieved robust transfection of neurons in the DCN and IC as well as reliable axonal trafficking of Chronos and ChrimsonR to synaptic terminals in the IC. Additionally, we demonstrated that this technique enables the measurement and analysis of postsynaptic events, including PSP amplitude, halfwidth, decay time, and receptor pharmacology. Our experience suggests that this approach can be readily adapted to perform functional circuit mapping experiments throughout the auditory brainstem and beyond.
Overall, the specificity of optogenetic circuit mapping provides a distinct advantage over electrical stimulation of axons. Viral transfections provide the ability to spatially and molecularly restrict the expression of channelrhodopsins to a targeted population of presynaptic neurons. In contrast, electrical stimulation cannot differentiate between axons originating from different presynaptic nuclei when those axons are intermingled, as is the case in most brain regions. Electrical stimulation can also initiate both orthodromic and antidromic spikes, further complicating matters when a distal stimulation site contains axons originating from neurons located near the recording site.
To ensure stable expression and good axonal trafficking of a channelrhodopsin, choosing the right viral vector and serotype is paramount. We found that stable expression of Chronos in IC neurons was achieved with a serotype 1 rAAV including a Chronos or ChrimsonR construct combined with a woodchuck hepatitis posttranscriptional regulatory element (WPRE) and bovine growth hormone (BGH) polyadenylation signal. rAAV serotype 5 failed to produce functional opsins in the IC, but was functional in the DCN (data not shown). The rigorous validation of injection coordinates for every experiment and ensuring that opsin expression is restricted to the targeted brain region is similarly important. A meaningful identification of inputs is only possible if expression of the opsin is carefully checked and documented for every experiment.
The rAAV1.ChrimsonR construct used in the above protocol yielded stable expression and good axonal trafficking of the protein, even in long range projections from the DCN to the IC (Figure 3). This makes ChrimsonR a suitable opsin for (long-range) circuit mapping. A red-shifted opsin like ChrimsonR can be useful if the experimental parameters require light penetration deep into the tissue, but the experimenter must be aware that all currently available red-shifted opsins show some cross-activation with blue light. Although some studies have argued that ChrimsonR and Chronos may be separately activated6,14,16, our data suggest that great care must be taken with this approach. A recent report details additional methods that should be used if attempting to separate red-shifted opsins from Chronos13. Therefore, when using ChrimsonR and Chronos in two color CRACM experiments, carefully designed control experiments need to be executed to ensure clear separation of blue and red-shifted opsin activation.

Synaptic connections are critical to neural circuit function, but the precise topology and physiology of synapses within neural circuits are often difficult to probe experimentally. This is because electrical stimulation, the traditional tool of cellular electrophysiology, indiscriminately activates axons near the stimulation site, and in most brain regions, axons from different sources (local, ascending, and/or descending) intertwine. However, by using channelrhodopsin assisted circuit mapping (CRACM)1,2, this limitation can now be overcome3. Channelrhodopsin (ChR2) is a light activated, cation-selective ion channel originally found in the green alga Chlamydomonas reinhardtii. ChR2 can be activated by blue light of a wavelength around 450-490 nm, depolarizing the cell through cation influx. ChR2 was first described and expressed in Xenopus oocytes by Nagel and colleagues4. Shortly after that, Boyden and colleagues5 expressed ChR2 in mammalian neurons and showed that they could use light pulses to reliably control spiking on a millisecond timescale, inducing action potentials ~10 ms after activation of ChR2 with blue light. Optogenetic channels with even faster kinetics have been found recently (e.g., Chronos6).
The basic approach to a CRACM experiment is to transfect a population of putative presynaptic neurons with a recombinant adeno-associated virus (rAAV) that carries the genetic information for a channelrhodopsin. Transfection of neurons with rAAV leads to the expression of the encoded channelrhodopsin. Typically, the channelrhodopsin is tagged with a fluorescent protein like GFP (Green Fluorescent Protein) or tdTomato (a red fluorescent protein), so that transfection of neurons in the target region can easily be confirmed with fluorescence imaging. Because rAAVs are non-pathogenic, have a low inflammatory potential and long-lasting gene expression7,8, they have become a standard technique to deliver channelrhodopsins to neurons. If, after transfection of a putative presynaptic population of neurons, activation of a channelrhodopsin through light flashes elicits postsynaptic potentials or currents in the target neurons, this is evidence of an axonal connection from the transfected nucleus to the recorded cell. Because severed axons in brain slice experiments can be driven to release neurotransmitter through channelrhodopsin activation, nuclei that lie outside of the acute slice but send axons into the postsynaptic brain region can be identified with CRACM. The power of this technique is that the connectivity and physiology of identified long range synaptic inputs can be directly investigated.
In addition to channelrhodopsins that are excitable by blue light, investigators have recently identified several red-shifted channelrhodopsins9,10, including Chrimson and its faster analog ChrimsonR, both of which are excited with red light of ~660 nm6. Red-shifted opsins are of interest because red light penetrates tissue better than blue light, and red light may have a lower cytotoxicity than blue light10,11,12. Red-shifted channelrhodopsins also open up the possibility of dual color CRACM experiments, where the convergence of axons from different nuclei on the same neuron can be tested in one experiment6,13,14. However, current red-shifted opsins often exhibit unwanted cross-activation with blue light15,16,17, making two color experiments difficult. In addition, some reports have indicated that ChrimsonR undergoes limited axonal trafficking, which can make it challenging to use ChrimsonR for CRACM experiments16,17.
Nearly all ascending projections from the lower auditory brainstem nuclei converge in the inferior colliculus (IC), the midbrain hub of the central auditory pathway. This includes projections from the cochlear nucleus (CN)18,19, most of the superior olivary complex (SOC)20, and the dorsal (DNLL) and ventral (VNLL) nuclei of the lateral lemniscus21. Additionally, a large descending projection from the auditory cortex terminates in the IC18,19,20,21,22, and IC neurons themselves synapse broadly within the local and contralateral lobes of the IC23. The intermingling of axons from many sources has made it difficult to probe IC circuits using electrical stimulation24. As a result, even though neurons in the IC perform computations important for sound localization and the identification of speech and other communication sounds25,26, the organization of neural circuits in the IC is largely unknown. We recently identified VIP neurons as the first molecularly identifiable neuron class in the IC27. VIP neurons are glutamatergic stellate neurons that project to several long-range targets, including the auditory thalamus and superior colliculus. We are now able to determine the sources and function of local and long-range inputs to VIP neurons and to determine how these circuit connections contribute to sound processing.
The protocol presented here is tailored to investigating synaptic inputs to VIP neurons in the IC of mice, specifically from the contralateral IC and the DCN (Figure 1). The protocol can be easily adapted to different sources of input, a different neuron type or a different brain region altogether. We also show that ChrimsonR is an effective red-shifted channelrhodopsin for long range circuit mapping in the auditory brainstem. However, we demonstrate that ChrimsonR is strongly activated by blue light, even at low intensities, and thus, to combine ChrimsonR with Chronos in two-color CRACM experiments, careful controls must be used to prevent cross-activation of ChrimsonR.

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long-range channelrhodopsin-assisted circuit mapping of inferior colliculus neurons with blue and red-shifted channelrhodopsins

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, making it difficult to isolate inputs from individual sources with electrical stimulation. However, by using channelrhodopsin assisted circuit mapping (CRACM), this limitation can now be overcome. Here, we report a method to use CRACM to map ascending inputs from lower auditory brainstem nuclei and commissural inputs to an identified class of neurons in the inferior colliculus (IC), the midbrain nucleus of the auditory system. In the IC, local, commissural, ascending, and descending axons are heavily intertwined and therefore indistinguishable with electrical stimulation. By injecting a viral construct to drive expression of a channelrhodopsin in a presynaptic nucleus, followed by patch clamp recording to characterize the presence and physiology of channelrhodopsin-expressing synaptic inputs, projections from a specific source to a specific population of IC neurons can be mapped with cell type-specific accuracy. We show that this approach works with both Chronos, a blue light-activated channelrhodopsin, and ChrimsonR, a red-shifted channelrhodopsin. In contrast to previous reports from the forebrain, we find that ChrimsonR is robustly trafficked down the axons of dorsal cochlear nucleus principal neurons, indicating that ChrimsonR may be a useful tool for CRACM experiments in the brainstem. The protocol presented here includes detailed descriptions of the intracranial virus injection surgery, including stereotaxic coordinates for targeting injections to the dorsal cochlear nucleus and IC of mice, and how to combine whole cell patch clamp recording with channelrhodopsin activation to investigate long-range projections to IC neurons. Although this protocol is tailored to characterizing auditory inputs to the IC, it can be easily adapted to investigate other long-range projections in the auditory brainstem and beyond.

We crossed VIP-IRES-Cre mice (Viptm1(cre)Zjh/J) and Ai14 Cre-reporter mice (B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J) to generate F1 offspring in which VIP neurons express the fluorescent protein tdTomato. F1 offspring of either sex were used, aged postnatal day (P) 21 to P70. A total of 22 animals were used in this study.
Stereotaxic injection of AAV1.Syn.Chronos-GFP.WPRE.bGH into the right IC of VIP-IRES-Cre x Ai14 mice using coordinates shown in Table 1 resulted in strong Chronos-EGFP expression in the right IC (Figure 2A). Visual inspection of Chronos-EGFP fluorescence indicated that most of the somata labeled in the right IC were located in the central nucleus of the IC (ICc), but labeled somata were sometimes also present in the dorsal cortex of the IC (ICd) and occasionally in the lateral cortex of the IC (IClc). The targeting and extent of transfection should be checked for every animal used in an experiment, as expression of channelrhodopsins in non-targeted regions can lead to false positives. To achieve broader or more restricted expression of Chronos, the amount of deposited virus as well as the stereotaxic coordinates can be easily adjusted to achieve the desired outcome.
Stereotaxic injection of AAV1.Syn.Chronos-GFP.WPRE.bGH into the DCN (see coordinates in Table 1) of VIP-IRES-Cre x Ai14 mice resulted in strong transfection of DCN neurons (Figure 2B). To confirm selective transfection of the DCN, the brainstem of every animal should be sliced to verify that EGFP expression was present and limited to the DCN. If there is no transfection or if there is considerable expression of EGFP in the auditory nerve or VCN, recordings should not be performed. Using the coordinates shown in Table 1 with a total injection volume of 40 nL, Chronos-EGFP expression will be limited to the DCN in most cases. EGFP-labeled axons were present in the left (contralateral) ICc 3 weeks after injection for both IC and DCN injection sites (Figure 2A right, 2B right).
To test the long-range trafficking of ChrimsonR, AAV1.Syn.ChrimsonR-tdTomato.WPRE.bGH was injected into the right DCN of VIP-IRES-Cre x Ai14 mice, using the same coordinates as with Chronos injections. ChrimsonR injection led to strong expression in the DCN, with tdTomato fluorescence visible in cells and fibers (Figure 3A). In the contralateral ICc, fibers strongly labeled with tdTomato were clearly visible after 3 weeks, demonstrating the long-range trafficking capability of the ChrimsonR-tdTomato construct when injected into auditory brainstem nuclei (Figure 3B). Optical activation of ChrimsonR elicited EPSPs in IC VIP neurons (Figure 3C), indicating that ChrimsonR is a useful tool for long-range CRACM experiments when the experimental parameters demand the use of red light instead of blue light. However, we found that ChrimsonR was readily activated with blue light, showing the same threshold for blue light activation as Chronos (Figure 4). The sensitivity of ChrimsonR to blue light means that special care must be taken to distinguish between inputs transfected with ChrimsonR or Chronos in the same animal13.
When targeting recordings to VIP neurons in the contralateral (left) ICc after injections into the right IC, blue light flashes elicited excitatory postsynaptic potentials (EPSPs) or inhibitory postsynaptic potentials (IPSPs). This confirms commissural projections to VIP neurons. To analyze commissural EPSPs and IPSPs separately, we used receptor antagonists to block IPSPs during EPSP recordings, and vice versa. Representative EPSPs and IPSPs recorded during CRACM experiments are shown in Figure 5. IPSPs were observed in 6 out of 12 tested ICc VIP neurons. IPSPs were small (1.53 mV &#177; 0.96 mV) and had moderate kinetics. The 10 - 90% rise times of IPSPs were 7.8 ms &#177; 2.1 ms, halfwidths were 15.1 ms &#177; 6.8 ms, and decay time constants were 32.4 ms &#177; 17.0 ms (Figure 5A, left). IPSPs were mediated by GABAA receptors, as they were blocked by 5 &#181;M gabazine, a GABAA receptor antagonist (Figure 5A, right; n = 6). EPSPs were observed in 11 out of 27 ICc VIP neurons tested. EPSPs were also small (1.52 mV &#177; 1.08 mV) and had moderate kinetics. The 10 - 90% rise times of EPSPs were 8.3 ms &#177; 4.3 ms, halfwidths were 19.6 ms &#177; 7.6 ms, and decay time constants were 43.5 ms &#177; 16.8 ms (Figure 5B &#34;Control&#34;). EPSPs were mediated by AMPA and NMDA receptors. Application of 50 &#181;M D-AP5, an NMDA receptor antagonist, reduced EPSP halfwidth (14.3 ms &#177; 4.7 ms, p = 0.006) and trended toward reducing the rise time (6.3 ms &#177; 1.6 ms, p = 0.09), decay time constant (30.6 ms &#177; 7.3 ms, p = 0.06), and EPSP amplitude (1.38 &#177; 0.33 mV, p = 0.105; ANOVA for repeated measurements with Tukey post-hoc test). Application of 10 &#181;M NBQX, an AMPA receptor antagonist, blocked the remainder of the EPSP (Figure 5B &#34;+ AP5 &#38; NBQX&#34;).
Recordings of VIP neurons in the left ICc after DCN injections revealed EPSPs evoked by blue light flashes, confirming synaptic inputs from the DCN to VIP neurons in the IC. DCN CRACM experiments were conducted with GABAergic and glycinergic blockers in the bath to block spontaneous IPSPs. We found that 2-5 ms pulses of blue light elicited EPSPs in 19 of 25 neurons tested. Light-evoked EPSPs had moderate amplitudes (2.85 mV &#177; 2.98 mV) and relatively slow rise times (4.2 ms &#177; 1.3 ms), halfwidths (20.6 ms &#177; 14.4 ms) and decay time constants (22.0 ms &#177; 6.7 ms) (n = 6 cells, data not shown).
All PSPs evoked by Chronos or ChrimsonR activation were elicited in an all-or-none fashion and did not scale their amplitude with increasing optical power (data not shown). However, this result depends on the number and physiology of the transfected synapses providing input to a particular neuron and is therefore likely to vary depending on the particular combination of axonal projection and neuron type being investigated.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60760/60760fig1.jpg" /> 
Figure 1: rAAV injection sites and experimental setup. (A) Injection site and experimental setup to investigate commissural projections. Left: An rAAV construct (e.g., rAAV1.Syn.Chronos-GFP.WPRE.bGH) is injected into the right IC and targeted recordings are performed in the contralateral IC. Right: During patch clamp recordings, Chronos-transfected fibers are excited with blue light to elicit postsynaptic potentials. (B) Injection site and experimental setup to investigate long-range projections from DCN to IC. Left: The ChrimsonR construct is injected into the right DCN and targeted recordings are performed in the contralateral IC. Right: During patch clamp recordings, ChrimsonR-transfected fibers are excited with red light to record ChrimsonR evoked postsynaptic potentials. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60760/60760fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60760/60760fig2a.jpg" /> 
Figure 2: Chronos-EGFP expression after injections into IC and DCN. (A) Left: Image of coronal IC slice showing tdTomato-labeled VIP neurons throughout the IC (magenta) and Chronos-EGFP expression in somata localized to the injection sites in the right IC (green). Right: Higher magnification image showing a recorded VIP neuron (white-green) in the IC contralateral to the injection site. Green puncta are Chronos-EGFP expressing projections from the contralateral IC. (B) Image of coronal brainstem slice showing Chronos-EGFP expression in the DCN after rAAV-Chronos-EGFP injection. Left: Image of the DCN, showing transfected DCN and EGFP-positive fibers entering the dorsal acoustic stria. Middle: Higher magnification image showing Chronos-transfected cell bodies in the DCN. Right: Chronos-EGFP expression in fibers and terminals in the contralateral IC from long range DCN-IC projections. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60760/60760fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60760/60760fig3a.jpg" /> 
Figure 3: ChrimsonR-tdTomato expression in the DCN and DCN projections to the IC. (A) Image of a coronal brainstem slice from a mouse in which the right DCN was injected with rAAV1.Syn.ChrimsonR-tdTomato.WPRE.bGH. Left: Image of ChrimsonR expression in the right DCN. Right: Higher magnification image of right DCN showing strong transfection of neurons with ChrimsonR. (B) High magnification image of ChrimsonR-transfected DCN axons in the IC. (C) Optogenetically evoked EPSPs recorded from a VIP neuron in the IC contralateral to the rAAV-ChrimsonR injected DCN. Original traces are shown in light grey and average EPSP is in red. Orange box indicates timing of 590 nm light pulse. Scale bars = 20 ms/0.5 mV. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60760/60760fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60760/60760fig4a.jpg" /> 
Figure 4: Activation of ChrimsonR by low levels of blue light. (A) Activation of Chronos in commissural projections by pulses of 470 nm blue light. Top: Original traces (light grey) and average (cyan) of Chronos-driven IPSPs, evoked at an optical power slightly above the threshold for Chronos activation (scale bars show 20 ms/0.5 mV). Bottom: Relationship between optical power at 470 nm and the probability of observing a Chronos-evoked PSP. (B) Activation of ChrimsonR with 470 nm blue light. Top: Original traces (light grey) and average (red) of ChrimsonR-driven EPSPs, evoked with 470 nm blue light at the same optical power used in A, top (scale bars show 20 ms/0.5 mV). Bottom: Relationship between optical power at 470 nm and the probability of observing a ChrimsonR-driven PSP. Note that the threshold for blue light activation of ChrimsonR PSPs was identical to the threshold for eliciting Chronos PSPs. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60760/60760fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60760/60760fig5a.jpg" /> 
Figure 5: Characterization of light-evoked PSPs from commissural synapses onto VIP neurons. The right IC was injected with rAAV1.Syn.Chronos-GFP.WPRE.bGH and recordings were made from VIP neurons in the contralateral ICc. (A) Optogenetically-evoked IPSPs were evoked by 2-5 ms blue light flashes (left) while EPSPs were blocked by 10 &#181;M NBQX and 50 &#181;M D-AP5. IPSPs were abolished by gabazine (right). (B) Optogenetically-evoked EPSPs were evoked by 2-5 ms blue light flashes (left) while IPSPs were blocked with 1 &#181;M strychnine and 5 &#181;M gabazine. Wash-in of 50 &#181;M D-AP5 significantly reduced the halfwidth and decay time constant of light-evoked EPSPs (middle). Wash-in of 10 &#181;M NBQX abolished the remaining EPSP (right). Original traces in A and B are shown in light grey, and averages from up to 50 individual traces are shown in black. From Goyer et al., 2019. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60760/60760fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>X (lateral)</td>
			<td>Y (caudal)</td>
			<td>Z (depth)</td>
		</tr>
		<tr>
			<td>Right IC penetration 1</td>
			<td>1,000 &#181;m</td>
			<td>-900 &#181;m</td>
			<td>2,250-1500 &#181;m 
			(250 &#181;m increments)</td>
		</tr>
		<tr>
			<td>Right IC penetration 2</td>
			<td>1,250 &#181;m</td>
			<td>-900 &#181;m</td>
			<td>2,250-1750 &#181;m 
			(250 &#181;m increments)</td>
		</tr>
		<tr>
			<td>Right DCN</td>
			<td>2,155 &#181;m</td>
			<td>-1325 &#181;m</td>
			<td>4,750 &#181;m, 4,550 &#181;m</td>
		</tr>
	</tbody>
</table>
Table 1: Stereotaxic coordinates for rAAV injections into IC and DCN. All coordinates are relative to lambda in &#181;m. Z coordinates are measured from the dorsal surface of the skull at lambda.

Watch this Scientific Journal Video about Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins at JoVE.com

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

Channelrhodopsin-assisted circuit mapping (CRACM) is a precision technique for functional mapping of long-range neuronal projections between anatomically and/or genetically identified groups of neurons. Here, we describe how to utilize CRACM to map auditory brainstem connections, including the use of a red-shifted opsin, ChrimsonR.

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, ...

long-range-channelrhodopsin-assisted-circuit-mapping-inferior

Nanyang Technological University

Research

JoVE Journal

Neuroscience

7.3K Views.  University of Michigan. Channelrhodopsin-assisted circuit mapping (CRACM) is a precision technique for functional mapping of long-range neuronal projections between anatomically and/or genetically identified groups of neurons. Here, we describe how to utilize CRACM to map auditory brainstem connections, including the use of a red-shifted opsin, ChrimsonR.

Video: Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.

This paper introduces an approach of combining laser scanning photostimulation with whole cell recordings in transgenic mice expressing GFP in limited inhibitory neuron populations. The technique allows for extensive mapping and quantitative analysis of local synaptic circuits of specific inhibitory cortical neurons.

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into ...

Using Affordable LED Arrays for Photo-Stimulation of Neurons

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single cells in response to electrical or pharmacological manipulations1,2. With the invention of methods to optically control genetically targeted neurons (optogenetics), researchers now have an unprecedented level of control over specific groups of neurons in the standard slice preparation. In particular, photosensitive channelrhodopsin-2 (ChR2) allows researchers to activate neurons with light3,4. By combining careful calibration of LED-based photostimulation of ChR2 with standard slice electrophysiology, we are able to probe with greater detail the role of adult-born interneurons in the olfactory bulb, the first central relay of the olfactory system. Using viral expression of ChR2-YFP specifically in adult-born neurons, we can selectively control young adult-born neurons in a milieu of older and mature neurons. Our optical control uses a simple and inexpensive LED system, and we show how this system can be calibrated to understand how much light is needed to evoke spiking activity in single neurons. Hence, brief flashes of blue light can remotely control the firing pattern of ChR2-transduced newborn cells.

Adult-born neurons expressing ChR2 can be manipulated in slice electrophysiological preparations in order to examine their contribution towards the function of olfactory neural circuits.

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single ...

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available to evoke excitatory junctional potentials (EJPs) in this preparation involves the use of suction electrodes. In both research and teaching labs, students often have difficulty maneuvering and manipulating this type of stimulating electrode. In the present work, we show how to remotely stimulate synaptic potentials at the larval NMJ without the use of suction electrodes. By expressing channelrhodopsin2 (ChR2) 4,5,6 in Drosophila motor neurons using the GAL4-UAS system 7, and making minor changes to a basic electrophysiology rig, we were able to reliably evoke EJPs with pulses of blue light. This technique could be of particular use in neurophysiology teaching labs where student rig practice time and resources are limited.

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

This procedure uses a blue light-activated algal channel and cell-specific genetic expression tools to evoke synaptic potentials with light pulses at the neuromuscular junction (NMJ) in Drosophila larvae. This technique is an inexpensive and easy-to-use alternative to suction electrode stimulation for synaptic physiology studies in research and teaching laboratories.

The Drosophila larval neuromuscular preparation has proven to be a useful tool for studying synaptic physiology1,2,3. Currently, the only means available ...

Biology

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

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

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

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

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits&#160;in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.

We describe a method for characterizing the functional topography and synaptic properties of forebrain circuits using an optogenetic approach to photostimulate neuronal populations&#160;in vitro.

The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to blindness in many retinal diseases. Calcium (Ca2+), a key second messenger in photoreceptor signaling and metabolism, has been proposed to be indirectly linked with photoreceptor degeneration in various animal models. Systematically studying these aspects of cone physiology and pathophysiology has been hampered by the difficulties of electrically recording from these small cells, in particular in the mouse where the retina is dominated by rod photoreceptors. To circumvent this issue, we established a two-photon Ca2+ imaging protocol using a transgenic mouse line that expresses the genetically encoded Ca2+ biosensor TN-XL exclusively in cones and can be crossbred with mouse models for photoreceptor degeneration. The protocol described here involves preparing vertical sections (&#8220;slices&#8221;) of retinas from mice and optical imaging of light stimulus-evoked changes in cone Ca2+ level. The protocol also allows &#8220;in-slice measurement&#8221; of absolute Ca2+ concentrations; as the recordings can be followed by calibration. This protocol enables studies into functional cone properties and is expected to contribute to the understanding of cone Ca2+ signaling as well as the potential involvement of Ca2+ in photoreceptor death and retinal degeneration.

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

We describe a protocol to monitor Ca2+ dynamics in the axon terminals of cone photoreceptors using an ex-vivo&#160;slice preparation of the mouse retina. This protocol allows comprehensive studies of cone Ca2+ signaling in an important mammalian model system, the mouse.

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. Traditional in vitro electrophysiological techniques use electrical stimulation to evoke synaptic transmission. A major drawback of this method is its nonspecific nature; all axons in the region of the stimulating electrode will be activated, making it difficult to attribute an effect to a particular afferent connection. This issue can be overcome by replacing electrical stimulation with optogenetic-based stimulation. We describe a method for combining optogenetics with in vitro patch-clamp recordings. This is a powerful tool for the study of both basal synaptic transmission and synaptic plasticity of precise anatomically defined synaptic connections and is applicable to almost any pathway in the brain. Here, we describe the preparation and handling of a viral vector encoding channelrhodopsin protein for surgical injection into a pre-synaptic region of interest (medial prefrontal cortex) in the rodent brain and making of acute slices of downstream target regions (lateral entorhinal cortex). A detailed procedure for combining patch-clamp recordings with synaptic activation by light stimulation to study short- and long-term synaptic plasticity is also presented. We discuss examples of experiments that achieve pathway- and cell-specificity by combining optogenetics and Cre-dependent cell labeling. Finally, histological confirmation of the pre-synaptic region of interest is described along with biocytin labeling of the post-synaptic cell, to allow further identification of the precise location and cell type.

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex

Here we present a protocol describing viral transduction of discrete brain regions with optogenetic constructs to permit synapse-specific electrophysiological characterization in acute rodent brain slices.

Studying the physiological properties of specific synapses in the brain, and how they undergo plastic changes, is a key challenge in modern neuroscience. ...

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

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Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

Optogenetic Stimulation of the Auditory Nerve

Imaging Ca²⁺ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Ex Vivo Optogenetic Interrogation of Long-Range Synaptic Transmission and Plasticity from Medial Prefrontal Cortex to Lateral Entorhinal Cortex