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 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.

<table><tbody><tr><td>AAV1.Syn.ChrimsonR-tdTomato.WPRE.bGH</td><td>Addgene</td><td>59171-AAV1</td><td></td></tr><tr><td>AAV1.Syn.Chronos-GFP.WPRE.bGH</td><td>Addgene</td><td>59170-AAV1</td><td></td></tr><tr><td>Ai14 reporter mice (B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)</td><td>Jackson Laboratory</td><td>stock #007914</td><td></td></tr><tr><td>Amber (590nm) LUXEON Rebel LED</td><td>Luxeon Star LEDs</td><td>SP-01-A8</td><td></td></tr><tr><td>Blue (470nm) LUXEON Rebel LED</td><td>Luxeon Star LEDs</td><td>SP-01-B4</td><td></td></tr><tr><td>Carproject (carprofen)</td><td>Henry Schein Animal Health</td><td>59149</td><td></td></tr><tr><td>Drummond glas capillaries</td><td>Drummond Scientific Company</td><td>3-000-203-G/X</td><td></td></tr><tr><td>Drummond Nanoject 3</td><td>Drummond Scientific Company</td><td>3-300-207</td><td></td></tr><tr><td>Electrode beveler</td><td>Sutter Instrument</td><td>FG-BV10-D</td><td></td></tr><tr><td>Ethilon 6-0 (0.8 metric) nylon sutures</td><td>Ethicon</td><td>local pharmacy</td><td></td></tr><tr><td>Fixed stage microscope</td><td>any</td><td>n/a</td><td></td></tr><tr><td>Gas anesthesia head holder</td><td>David Kopf Instruments</td><td>933-B</td><td></td></tr><tr><td>General surgery tools</td><td>Fine Science Tools</td><td>N/A</td><td></td></tr><tr><td>Golden A5 pet clipper</td><td>Oster</td><td>078005-010-003</td><td></td></tr><tr><td>Heating pad</td><td>Custom build</td><td>N/A</td><td></td></tr><tr><td>Hooded induction chamber w/ vacuum system</td><td>Patterson Scientific</td><td>78917760</td><td></td></tr><tr><td>Hot bead sterilizer Steri 250</td><td>Inotech</td><td>IS-250</td><td></td></tr><tr><td>Iodine solution 10%</td><td>MedChoice</td><td>local pharmacy</td><td></td></tr><tr><td>Isoflurane vaporizer</td><td>Patterson Scientific</td><td>07-8703592</td><td></td></tr><tr><td>Lidocain topical jelly 2%</td><td>Akorn</td><td>local pharmacy</td><td></td></tr><tr><td>Micro motor drill 1050</td><td>Henry Schein Animal Health</td><td>7094351</td><td></td></tr><tr><td>Micro motor drill bits 0.5 mm</td><td>Fine Science Tools</td><td>19007-05</td><td></td></tr><tr><td>Motorized Micromanipulator</td><td>Sutter Instrument</td><td>MP-285/R</td><td></td></tr><tr><td>Ophthalmic ointment Artificial Tears</td><td>Akorn</td><td>local pharmacy</td><td></td></tr><tr><td>P-1000 electrode puller</td><td>Sutter Instrument</td><td>P-1000</td><td></td></tr><tr><td>Patch clamp amplifier incl data acquisition software</td><td>any</td><td>n/a</td><td></td></tr><tr><td>Portable anethesia machine</td><td>Patterson Scientific</td><td>07-8914724</td><td></td></tr><tr><td>Small animal steroetaxic frame</td><td>David Kopf Instruments</td><td>930-B</td><td></td></tr><tr><td>Standard chemicals</td><td>local vendors</td><td>N/A</td><td></td></tr><tr><td>standard imaging solutions</td><td></td><td></td><td></td></tr><tr><td>Sterile towel drapes</td><td>Dynarex</td><td>4410</td><td></td></tr><tr><td>Surgical marker</td><td>Fine Science Tools</td><td>18000-30</td><td></td></tr><tr><td>Temperature controller</td><td>Custom build</td><td>N/A</td><td></td></tr><tr><td>Vibratome</td><td>any</td><td>n/a</td><td></td></tr><tr><td>VIP-IRES-Cre mice (Viptm1(cre)Zjh/J)</td><td>Jackson Laboratory</td><td>stock #010908</td><td></td></tr><tr><td>Water bath</td><td>any</td><td>n/a</td><td></td></tr></tbody></table>

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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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

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are reprogrammed from human fibroblasts using episomal vectors. Colonies of iPSCs can be observed 30 days after initiation of fibroblast reprogramming. Pluripotent colonies are manually picked and grown in neural induction medium to permit differentiation into neural progenitor cells (NPCs). iPSCs rapidly convert into neuroepithelial cells within 1 week and retain the capability to self-renew when maintained at a high culture density. Primary mouse NPCs are differentiated into astrocytes by exposure to a serum-containing medium for 7 days and form a monolayer upon which embryonic day 18 (E18) rat cortical neurons (transfected with channelrhodopsin-2 (ChR2)) are added. Human NPCs tagged with the fluorescent protein, tandem dimer Tomato (tdTomato), are then seeded onto the astrocyte/cortical neuron culture the following day and allowed to differentiate for 28 to 35 days. We demonstrate that this system forms synaptic connections between iPSC-derived neurons and cortical neurons, evident from an increase in the frequency of synaptic currents upon photostimulation of the cortical neurons. This co-culture system provides a novel platform for evaluating the ability of iPSC-derived neurons to create synaptic connections with other neuronal populations.

A protocol to generate a co-culture system consisting of neurons derived from induced pluripotent stem cells (iPSCs), primary cortical neurons and astrocytes is described. This co-culture system allows detection of the formation of synaptic contacts and circuits between new, iPSC-derived neurons and pre-existing cortical neurons expressing channelrhodopsin-2.

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are ...

Developmental Biology

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 ...

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, ...

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 ...

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic network, electrophysiological recordings are frequently used to study connections among individual neurons. We have optimized a flat-mount preparation for patch clamp recording of genetically marked neurons in both GCL (ganglion cell layer) and INL (inner nuclear layer) of mouse retinas. Recording INL neurons in flat-mounts is favored over slices because both vertical and lateral connections are preserved in the former configuration, allowing retinal circuits with large lateral components to be studied. We have used this procedure to compare responses of mirror-partnered neurons in retinas such as the cholinergic starburst amacrine cells (SACs).

This protocol demonstrates how to perform whole-cell patch clamp recording on retinal neurons from a flat-mount preparation.

The mammalian retina is a layered tissue composed of multiple neuronal types. To understand how visual signals are processed within its intricate synaptic ...

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the identification of the neuronal type in in vivo electrophysiological experiments in broad brain regions. In optogenetics experiments, it is critical to deliver the light to the recording site. However, it is often hard to deliver the stimulation light to the deep brain regions from the brain&#39;s surface. Especially, it is difficult for the stimulation light to reach the deep brain regions when the optical transparency of the brain surface is low, as is often the case with recordings from awake animals. Here, we describe a method to record spike responses to the light from an awake mouse using a custom-made glass optrode. In this method, the light is delivered through the recording glass electrode so that it is possible to reliably stimulate the recorded neuron with light in the deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the ...

Two-Photon and Wide-Field Calcium Imaging in the Superior Colliculus

Calcium Imaging in Mouse Superior Colliculus

The superior colliculus (SC), an evolutionarily conserved midbrain structure in all vertebrates, is the most sophisticated visual center before the emergence of the cerebral cortex. It receives direct inputs from ~30 types of retinal ganglion cells (RGCs), with each encoding a specific visual feature. It remains elusive whether the SC simply inherits retinal features or if additional and potentially de novo processing occurs in the SC. To reveal the neural coding of visual information in the SC, we provide here a detailed protocol to optically record visual responses with two complementary methods in awake mice. One method uses two-photon microscopy to image calcium activity at single-cell resolution without ablating the overlaying cortex, while the other uses wide-field microscopy to image the whole SC of a mutant mouse whose cortex is largely undeveloped. This protocol details these two methods, including animal preparation, viral injection, headplate implantation, plug implantation, data acquisition, and data analysis. The representative results show that the two-photon calcium imaging reveals visually evoked neuronal responses at single-cell resolution, and the wide-field calcium imaging reveals&#160;neural activity across the entire SC. By combining these two methods, one can reveal the neural coding in the SC at different scales, and such combination can also be applied to other brain regions.

Author Spotlight: Unveiling Neural Coding and Mechanisms of Visual Processing in the Superior Colliculus 

This protocol details the procedure for imaging calcium responses in the superior colliculus (SC) of awake mice, including imaging single-neuron activity with two-photon microscopy while leaving the cortex intact in wild-type mice, and imaging the entire SC with wide-field microscopy in partial-cortex mutant mice.

The superior colliculus (SC), an evolutionarily conserved midbrain structure in all vertebrates, is the most sophisticated visual center before the ...

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

Summary

Explore More Videos

Chapters in this video

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

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

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

Patch Clamp Recording of Starburst Amacrine Cells in a Flat-mount Preparation of Deafferentated Mouse Retina

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

Calcium Imaging in Mouse Superior Colliculus