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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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, ...

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

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

Functional Mapping with Simultaneous MEG and EEG

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the processing of simple sensory stimuli. We will use somatosensory stimuli to locate the hand somatosensory areas, auditory stimuli to locate the auditory cortices, visual stimuli in four quadrants of the visual field to locate the early visual areas. These type of experiments are used for functional mapping in epileptic and brain tumor patients to locate eloquent cortices. In basic neuroscience similar experimental protocols are used to study the orchestration of cortical activity. The acquisition protocol includes quality assurance procedures, subject preparation for the combined MEG/EEG study, and acquisition of evoked-response data with somatosensory, auditory, and visual stimuli. We also demonstrate analysis of the data using the equivalent current dipole model and cortically-constrained minimum-norm estimates. Anatomical MRI data are employed in the analysis for visualization and for deriving boundaries of tissue boundaries for forward modeling and cortical location and orientation constraints for the minimum-norm estimates.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to map brain areas involved in the processing of simple sensory stimuli.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the ...

Revealing Neural Circuit Topography in Multi-Color

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably label the global patterning of multiple topographic projections. The cerebellum is an ideal model to study the orderly arrangement of neural circuits. For example, the compartmental organization of spinocerebellar mossy fibers has proven to be an indispensable system for studying mossy fiber patterning. We recently showed that wheat germ agglutinin (WGA) conjugated to Alexa 555 and 488 can be used for tracing spinocerebellar mossy fiber projections in developing and adult mice (Reeber et al. 2011). We found three major properties that make the WGA-Alexa tracers desirable tools for labeling neural projections. First, Alexa fluorophores are intense and their brightness allows for wholemount imaging directly after tracing. Second, WGA-Alexa tracers label the entire trajectory of developing and adult neural projections. Third, WGA-Alexa tracers are rapidly transported in both retrograde and anterograde directions. Here, we describe in detail how to prepare the tracers and other required tools, how to perform the surgery for spinocerebellar tracing and how best to image traced projections in three dimensions. In summary, we provide a step-by-step tracing protocol that will be useful for deciphering the organization and connectivity of functional maps not only in the cerebellum but also in the cortex, brainstem, and spinal cord.

We provide a practical guide for delivering tracers in vivo and use the spinocerebellar pathway as a model system to demonstrate essential steps for successful neuronal circuit analysis in mice. We describe in detail our versatile tracing protocol that exploits wheat germ agglutinin (WGA) conjugated to Alexa fluorophores.

Neural circuits are organized into functional topographic maps. In order to visualize complex circuit architecture we developed an approach to reliably ...

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of neurophysiology, neuroethology, and developmental biology. The goal of this report is to consolidate experimental techniques from the leech system into a single article that will be of use to physiologists with expertise in other nervous system preparations, or to biology students with little or no electrophysiology experience. We demonstrate how to dissect the leech for recording intracellularly from identified neural circuits in the ganglion. Next we show how individual cells of known function can be removed from the ganglion to be cultured in a Petri dish, and how to record from those neurons in culture. Then we demonstrate how to prepare a patch of innervated skin to be used for mapping sensory or motor fields. These leech preparations are still widely used to address basic electrical properties of neural networks, behavior, synaptogenesis, and development. They are also an appropriate training module for neuroscience or physiology teaching laboratories.

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

This article describes three nervous system preparations using leeches: intracellular recording from neurons in ventral ganglia, culturing neurons from ventral ganglia, and recording from a patch of innervated skin to map sensory fields.

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma establishes a lifelong persistent infection in the brain. While this brain infection is asymptomatic in most immunocompetent people, in the developing fetus or immunocompromised individuals such as acquired immune deficiency syndrome (AIDS) patients, this predilection for and persistence in the brain can lead to devastating neurologic disease. Thus, it is clear that the brain-Toxoplasma interaction is critical to the symptomatic disease produced by Toxoplasma, yet we have little understanding of the cellular or molecular interaction between cells of the central nervous system (CNS) and the parasite. In the mouse model of CNS toxoplasmosis it has been known for over 30 years that neurons are the cells in which the parasite persists, but little information is available about which part of the neuron is generally infected (soma, dendrite, axon) and if this cellular relationship changes between strains. In part, this lack is secondary to the difficulty of imaging and visualizing whole infected neurons from an animal. Such images would typically require serial sectioning and stitching of tissue imaged by electron microscopy or confocal microscopy after immunostaining. By combining several techniques, the method described here enables the use of thick sections (160 &#181;m) to identify and image whole cells that contain cysts, allowing three-dimensional visualization and analysis of individual, chronically infected neurons without the need for immunostaining, electron microscopy, or serial sectioning and stitching. Using this technique, we can begin to understand the cellular relationship between the parasite and the infected neuron.

3-D Imaging and Analysis of Neurons Infected In Vivo with Toxoplasma gondii

Using this protocol, we were able to image 160 &#181;m thick brain sections from mice infected with the parasite Toxoplasma gondii, which enables visualization and analysis of the spatial relationship between the encysting parasite and the infected neuron.

Toxoplasma gondii is an obligate, intracellular parasite with a broad host range, including humans and rodents. In both humans and rodents, Toxoplasma ...

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 ...

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 ...

Translational Brain Mapping at the University of Rochester Medical Center: Preserving the Mind Through Personalized Brain Mapping

The Translational Brain Mapping Program at the University of Rochester is an interdisciplinary effort that integrates cognitive science, neurophysiology, neuroanesthesia, and neurosurgery. Patients who have tumors or epileptogenic tissue in eloquent brain areas are studied preoperatively with functional and structural MRI, and intraoperatively with direct electrical stimulation mapping. Post-operative neural and cognitive outcome measures fuel basic science studies about the factors that mediate good versus poor outcome after surgery, and how brain mapping can be further optimized to ensure the best outcome for future patients. In this article, we describe the interdisciplinary workflow that allows our team to meet the synergistic goals of optimizing patient outcome and advancing scientific understanding of the human brain.

This article provides an overview of a multi-modal brain mapping program designed to identify regions of the brain that support critical cognitive functions in individual neurosurgery patients.

The Translational Brain Mapping Program at the University of Rochester is an interdisciplinary effort that integrates cognitive science, neurophysiology, ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Meta-analysis of Voxel-Based Neuroimaging Studies using Seed-based d Mapping with Permutation of Subject Images (SDM-PSI)

Most methods for conducting meta-analysis of voxel-based neuroimaging studies do not assess whether effects are not null, but whether there is a convergence of peaks of statistical significance, and reduce the assessment of the evidence to a binary classification exclusively based on p-values (i.e., voxels can only be &ldquo;statistically significant&rdquo; or &ldquo;non-statistically significant&rdquo;). Here, we detail how to conduct a meta-analysis using Seed-based d Mapping with Permutation of Subject Images (SDM-PSI), a novel method that uses a standard permutation test to assess whether effects are not null. We also show how to grade the strength of the evidence according to a set of criteria that considers a range of statistical significance levels (from more liberal to more conservative), the amount of data or the detection of potential biases (e.g., small-study effect and excess of significance). To exemplify the procedure, we detail the conduction of a meta-analysis of voxel-based morphometry studies in obsessive-compulsive disorder, and we provide all the data already extracted from the manuscripts to allow the reader to replicate the meta-analysis easily. SDM-PSI can also be used for meta-analyses of functional magnetic resonance imaging, diffusion tensor imaging, position emission tomography and surface-based morphometry studies.

Meta-analysis of Voxel-Based Neuroimaging Studies using Seed-based d Mapping with Permutation of Subject Images SDM-PSI

We detail how to conduct a meta-analysis of voxel-based neuroimaging studies using Seed-based d Mapping with Permutation of Subject Images (SDM-PSI).