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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<li>Oda, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Crystal+structure+of+the+red+light-activated+channelrhodopsin+Chrimson%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Crystal structure of the red light-activated channelrhodopsin Chrimson.</a> Nature Communications. 9, (2018).</li>
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</ol>

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 e...

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 r...

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, ...

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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 ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

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

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

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

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

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

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

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

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

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

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 ...

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 ...

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 ...

Optogenetic Activation of Afferent Pathways in Brain Slices and Modulation of Responses by Volatile Anesthetics

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect distinct cellular and network components of these circuits remains unclear. Ex vivo brain slices provide a means by which investigators may probe discrete components of complex networks and disentangle potential mechanisms underlying the effects of volatile anesthetics on evoked responses. To isolate potential cell type- and pathway-specific drug effects in brain slices, investigators must be able to independently activate afferent fiber pathways, identify non-overlapping populations of cells, and apply volatile anesthetics to the tissue in aqueous solution. In this protocol, methods to measure optogenetically-evoked responses to two independent afferent pathways to neocortex in ex vivo brain slices are described. Extracellular responses are recorded to assay network activity and targeted whole-cell patch clamp recordings are conducted in somatostatin- and parvalbumin-positive interneurons. Delivery of physiologically relevant concentrations of isoflurane via artificial cerebral spinal fluid to modulate cellular and network responses is described.

Ex vivo brain slices can be used to study the effects of volatile anesthetics on evoked responses to afferent inputs. Optogenetics are employed to independently activate thalamocortical and corticocortical afferents to non-primary neocortex, and synaptic and network responses are modulated with isoflurane.

Anesthetics influence consciousness in part via their actions on thalamocortical circuits. However, the extent to which volatile anesthetics affect ...

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

Summary

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

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

Laser-scanning Photostimulation of Optogenetically Targeted Forebrain Circuits

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