This work was supported by grants from the National Eye Institute R01EY017925 and R21EY020985 and funding from the Dana &amp; Whitehall Foundations to P.K. We also thank Matthew Petrella for assistance with surgical procedures; Grace Dion for tracing the dendrites shown in Figure 5A; and Pratik Chhatbar for comments on the manuscript.

1. Surgical Preparation
<ol>
	<li>Induce anesthesia and continuously monitor heart rate, end tidal CO2, EEG, and temperature. All procedures were approved by the Institutional Animal Care and Use Committee of the Medical University of South Carolina and were based on those we previously published9,15.</li>
	<li>Expose the dorsal surface of the skull by cutting the skin with a scalpel blade. Dissect the connective tissues overlying the bone using a Brudon curette. Clean the bone using cotton tipped applicators and cotton gauze. Apply bone wax (as needed) to the skull, to stop occasional bleeding through small emissary veins.</li>
	<li>Attach a titanium or stainless steel head plate to the skull (over the region of interest where the craniotomy will be performed) by using dental cement mixed with black paint (see Materials). A rectangular opening in the center of the head plate provides access for the craniotomy and optical imaging. Attach a metal chamber on top of the head plate. This chamber will serve as a fluid reservoir for imaging with the water immersion objective lens.</li>
	<li>Secure the head plate to an xy stage using metal posts.</li>
	<li>Thin a large area of the bone within the rectangular opening of the headplate by using a dental drill. The area to be thinned must extend well beyond the boundaries of the planned craniotomy. The thick skull of non-rodent mammals must be thinned if high-resolution images are to be acquired from the neocortex because at the craniotomy site, the thickness of bone will dictate the thickness of the agarose between the coverglass and neocortical surface (see below).</li>
	<li>Drill a 2 x 2 mm square outline in the bone for the craniotomy. Rinse the chamber periodically with fresh artificial cerebrospinal fluid (ACSF) to remove bone shards and minimize heat dissipation to the underlying cortex. Apply bone wax as needed, to stop occasional bleeding.</li>
	<li>Lift the bone flap with a #3 forceps.</li>
	<li>Remove all but the bottom layer of dura using a fine forceps (#5CO) and Vannas spring scissors. The dura in non-rodent mammals is thick and opaque but can be removed in distinct layers. Bleeding is rare but can be stopped with Gelfoam and any blood residue should be gently removed from the remaining layer of dura using forceps and rinsing with ACSF. The final layer of dura is transparent and pial vessels should be clearly visible through it. Leaving this final layer of dura intact during intrinsic signal imaging will increase the success rate of bulk loading of fluorescent calcium indicators for the subsequent two-photon imaging phase of the experiment (see Discussion).</li>
	<li>Apply a drop of warm 2% agarose (dissolved in ACSF) and immediately place a coverglass over the craniotomy. Check the craniotomy to ensure that respiratory and cardiovascular pulsations are minimal by using a surgical dissecting microscope or the oculars of the two-photon microscope in bright-field mode.</li>
</ol>
2. Intrinsic Signal Optical Imaging
<ol>
	<li>Add gel-foam soaked with ACSF around the outside of the coverglass to prevent the agarose from drying during intrinsic signal imaging.</li>
	<li>Attach the intrinsic signal imaging CCD camera to the standard C-mount port on top of the two-photon microscope (Figure 1). Place the illuminating light source on the air table near the xy stage (Figure 1A). Position and secure the light guides over the craniotomy and headplate/chamber (Figure 1B). Retract the primary dichroic and the mirror below the C-mount port so that the reflected red light needed for intrinsic signals will pass directly from the craniotomy to the CCD camera (Figure 1C-D). Using the 4x air objective (0.13 NA, 17 mm WD), a 2 x 2 mm field of view is available for intrinsic signal imaging.</li>
	<li>Insert a heat filter in the light path to prevent the cortex from heating. Use a filter that passes green light (546 nm center &lambda;) and record a digital reference image of the cortical surface blood vessels.</li>
	<li>Move the z focus to 500 &mu;m below the cortical surface. Use a red filter (630 nm center &lambda;) to illuminate the cortex for measuring intrinsic signals. Position the light guides such that the cortical surface is uniformly illuminated. Shield the craniotomy from the light produced by the visual stimulus display. Present a sequence of visual stimuli and record reflectance data images. To generate an orientation map within 10 min, collect data corresponding to 4 repetitions of a sequence of 8 stimuli (4 orientations and two directions for each orientation).</li>
	<li>Analyze the intrinsic signal data to generate a false color orientation map. Select potential target regions for two-photon labeling of neurons, e.g., orientation pinwheel singularities - points in the map where all preferred orientations converge (Figure 2A). Overlay the orientation map and the image of the cortical surface vasculature (Figure 2B-C). Use the layout of functional domains and the location of large surface vessels to select a target, avoiding locations with large blood vessels and arterioles15.</li>
</ol>
3. Bulk Loading of Fluorescent Calcium Indicators
<ol>
	<li>Prepare a solution of 20% Pluronic/DMSO by adding 1 g Pluronic in 5 &mu;l DMSO. Heat the mixture at 60 &deg;C for 10 min to completely dissolve the Pluronic. Allow the mixture to cool to room temperature. Add 4 &mu;l of the Pluronic/DMSO mixture to a 50 &mu;g vial of the fluorescent calcium indicator Oregon Green 488 Bapta-1 AM (OGB-1 AM). Then add 1 &mu;l of a red dye (Alexa Fluor 594, 2 mM stock solution) and 35 &mu;l of pipette solution (see Materials) for a final concentration of 1 mM OGB-1 AM. Sonicate the solution for &frac12; hr in a chilled water bath and then centrifuge with a 0.45 &mu;m filter to remove impurities.</li>
	<li>Pull a long taper pipette with an outer tip diameter of 2.0-2.5 &mu;m. Load 5 &mu;l of the dye mixture into the pipette.</li>
	<li>Remove the final layer of dura to facilitate pipette insertion and obtain the highest quality two-photon images.</li>
	<li>Determine the exact location on the cortical surface that the pipette must be placed in order to reach its target in cortical layer 2/3. Our predetermined cortical target is a pinwheel singularity in the orientation map (Figure 2). The pipette must be driven into the cortex at a 30&deg; angle to pass between the chamber wall and the objective. Thus, to label the target region 200 &mu;m below the cortical surface, the pipette entry position on the cortical surface will be approximately 350 &mu;m lateral to the target position (Figure 3 A-B).</li>
	<li>Move the xy stage (on which the animal and the pipette micromanipulator are placed) to center the cortical surface entry position under the objective (Figure 3C). Use a 20x or 40x objective with a working distance (WD) of at least 3 mm (see Materials) to ensure that the pipette will easily enter the cortex without touching the objective or the bone along the wall of the craniotomy. Once the entry position is centered under the objective, raise the Z position of the objective by 2 mm (Figure 3D). With the green epi-fluorescence light turned on to visualize the red Alexa dye in the pipette, use the diagonal axis of the micromanipulator to move the pipette until its tip is centered (and focused) under the objective directly above the cortical surface entry position (Figure 3E).</li>
	<li>Lower the pipette vertically (along the z axis) until the pipette reaches the cortical surface (Figure 3F), while continuously viewing the pipette tip through the oculars of the microscope. Using bright-field illumination, the cortical surface blood vessels and remaining meningeal membranes (pia and arachnoid) are easily visible as the pipette approaches the cortical surface. If the pipette does not reach the desired entry position, raise the pipette and recalibrate targeting based on the vascular landmarks on the cortical surface.</li>
	<li>Remove the objective, wick excess cerebrospinal fluid from the craniotomy and dry the adjacent bone using lint-free absorption spears. Apply a drop of warm 3% agarose (dissolved in ACSF) to stabilize the cortex that is visible through the craniotomy (Figure 3G). In non-rodents, 3% agarose is necessary to dampen pulsations for successful uptake of fluorescent dyes by neurons. For intrinsic signal optical imaging only 2% agarose is needed because the combination of 2% agarose and the coverglass provides the necessary stability. Here, no coverglass is used for the dye-loading step so 3% agarose is necessary. After the agarose has solidified, insert a 40x objective (0.8 NA, 3.3 mm WD) and re-fill the chamber with ACSF.</li>
	<li>Switch from bright-field to two-photon viewing and locate the pipette tip. Using the diagonal axis of the micromanipulator, move the pipette until its tip reaches the desired depth in the cortex (Figure 3H). Occasional pressure ejection puffs of the dye mixture (a few psi, each pulse 0.1 sec) will help prevent the pipette tip from clogging. Because OGB-1 AM only becomes visible upon entering cells, the red Alexa dye is critical for visualization of the pipette tip inside the cortex.</li>
	<li>Before attempting to load neurons in cortical layer 2/3 with the full dose of dye, re-focus the objective on the cortical surface to confirm the depth of the pipette tip and ensure that the tip is in the target location based on the layout of surface blood vessels. If the pipette is in layer 2/3, small puffs of the dye mixture should illuminate the extracellular space and reveal a dense assembly of circular cell bodies as dark shadows.</li>
	<li>For loading neuronal cell bodies in a sphere of cortex 300-600 &mu;m diameter, inject the dye mixture into the extracellular space using 30-90 pulses, each pulse 1.0 sec, 2-10 psi. The pulses of pressure should not be so strong that the tissue moves. After the injection is complete, wait a few min, then remove the pipette and objective. Wait for 1 hr to allow the OGB-1 AM to be fully taken-up by neurons.</li>
	<li>Remove the agarose used for the dye loading and rinse the recording chamber. Put a small drop of 2-3% agarose on the surface and quickly place a coverglass over the craniotomy. Because the fluorescent dyes are light-sensitive, avoid over-exposing the cortex to bright-field or epi-fluorescent light. Insert a high NA (1.0) 20x objective into the objective holder and re-center the labeled cortical region under the objective using the xy stage. Shield the craniotomy from extraneous light sources, e.g., visual stimulus display, using multiple layers of blackout material.</li>
</ol>
4. Single-cell Electroporation of Fluorescent Dyes
<ol>
	<li>For studying dendritic morphology, prepare a solution of 100 &mu;M Alexa Fluor 594 (1:20 dilution in ACSF of 2 mM Alexa Fluor 594 stock). For functional imaging of dendrites an OGB salt may be used16,17.</li>
	<li>Pull a long taper pipette with a tip size of 1 &mu;m and load 5 &mu;l of dye into the pipette. All other steps of targeting a functional domain are identical to the bulk loading method (Figures 1-3). To position the pipette tip adjacent to a neuronal cell body membrane, monitor the increase in electrode impedance and apply pressure puffs of dye to illuminate the extracellular space and see cell bodies as shadows during two-photon imaging.</li>
	<li>When the pipette tip is adjacent to a neuronal cell body membrane, use the Axoporator 800A to apply 1-3 pulses (-8 to -10 V, 10 msec pulse). The immediate filling of the neuron with dye is readily visualized with two-photon microscopy. Remove the pipette and collect high-resolution images of dendrites and axons in vivo.</li>
</ol>

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No conflicts of interest declared.

We present a method to target the labeling of neuronal cell bodies (or dendrites and axons) in pre-determined functional micro-domains of the neocortex. Merging intrinsic signal optical imaging with two-photon microscopy offers the possibility of determining which synapses and cells contribute to the micro-domain of any functional map, whether neuronal selectivity correlates with the location of the neuron in a functional map, and the neuronal circuit components that change with visual experience7 or the application of clinically therapeutic drugs18.
No maps for stimulus orientation, direction, ocular dominance, or binocular disparity have been reported in the visual cortex of adult wild-type rodents6,19-21. However, the ease of loading neurons with fluorescent dyes and attaining imaging stability, together with the powerful genetic tools available for mice has resulted in the vast majority of studies using two-photon imaging of the cerebral cortex being performed in rodents instead of ferrets, cats, and monkeys.
High-resolution two-photon imaging of neurons in non-rodent mammals can be very challenging because of long surgery times, thicker bone and dura, and extra steps needed to keep respiratory and cardiovascular mediated pulsations of the brain to a minimum15. Transitions from surgery to intrinsic signal imaging and intrinsic to two-photon imaging need to be efficient, requiring timely preparation of reagents, dyes and the execution of micro-surgical procedures. Using the two-photon microscope for intrinsic signal imaging obviates the need for much of the equipment used in conventional intrinsic signal optical imaging rigs22 and reduces the time required to setup intrinsic imaging and transition to subsequent two-photon imaging.
Our approach is specifically designed to provide high-resolution two-photon imaging in pre-determined functional domains. This provides minimal cross talk of functional signals between neuronal cell bodies, and individual dendritic spines and axons can be resolved in vivo. Using a high numerical aperture (NA = 1.0) objective and filling the back aperture of the objective with beam expansion optics (see Supplementary Fig. 20 in Shen et al., 201215) are critical for high-resolution imaging. Depending on the speed of imaging in XY and/or Z, respiratory and cardiovascular pulsations may reduce the effective imaging resolution no matter how good the NA and optical light path are. By limiting the craniotomy size in non-rodents to 2 x 2 mm, using the relatively higher concentration of agarose with a coverglass, and performing a pneumothorax and lumbar suspension when necessary15, brain pulsations are limited to 1-2 &mu;m. Minimizing respiratory and cardiovascular induced pulsations of the neocortex to 1-2 &mu;m during bulk loading of calcium indicators also ensures that the cortex forms a tight seal around the inserted pipette and that no dye tracks up along the shank of pipette to the cortical surface. Selecting dye injection and imaging sites away from large arterioles is also critical for high quality functional imaging because relatively fast sensory-evoked hyperemia (hemodynamic) artifacts will be excluded15.
At the craniotomy site, leaving the final layer of dura intact throughout the intrinsic signal imaging and fluorescent dye preparation phases - thus removing it just before the pipette insertion into the cortex for dye loading - increases the success rate and quality of neuronal labeling in the following ways: (1) keeps the cortical surface clean and/or prevents dural tissue regrowth so that the pipette tip is less likely to get clogged and the transparency of the tissue is maintained, (2) avoids having the agarose used for intrinsic imaging touch the cortical surface blood vessels which may get damaged when the agarose is removed, (3) prevents the agarose used during intrinsic imaging from entering between the dura and cortex underneath the skull around the edges of the craniotomy. Agarose in these regions will increase the distance between the coverglass and the cortical surface which makes the insertion of the pipette more difficult, leads to compression of the cortical tissue and reduces the transparency for imaging.
When preparing the AM fluorescent dyes for bulk loading, the 20% Pluronic/DMSO mixture should never be used beyond 1 week from its initial preparation - old Pluronic/DMSO quenches fluorescence. For optimal bulk loading of many adjacent cell bodies with a functional indicator, e.g., OGB-1 AM, the dye mixture is stored on ice, loaded into the pipette as needed but always injected into the brain within 2 hr of its preparation.
During the bulk loading phase of the experiment, repeated 1 sec pulses of pressure rather than a continuous injection of 1 min allows for iterative calibration of ejection parameters, which ensures that excessive dye or pressure is not applied. Because of the potential tissue damage from multiple pipette insertions needed to label very large regions of tissue and potential toxicity from large volumes of Pluronic/DMSO from such a series of many injections, we prefer one 300-600 &mu;m dye injection site per 2 x 2 mm craniotomy. Compared to blind injections14, two-photon visually guided navigation around blood vessels, use of negative contrast of neuronal cell bodies to determine lamina position, and gauging the spread of the dye upon pressure ejection as an index of the volume of cells that will be labeled with OGB-1 AM, improves the quality of dye loading in non-rodent preparations.
The protocol we describe here leads to successful labeling of neurons with fluorescent dyes in targeted micro-domains in every animal attempted. For bulk loading with OGB-1 AM, we consistently obtain visually-evoked responses in 75-100% of simultaneously imaged neurons9,15 and the technique is sensitive to the detection of single action potentials in vivo23,24.

<table><tbody><tr><td colspan="4">&lt;strong&gt;1. Life support/experiment prep&lt;/strong&gt;</td></tr><tr><td>Isoflurane</td><td>Webster Vet</td><td>NDC 57319-474-05</td><td></td></tr><tr><td>Isoflurane vaporizer</td><td>Midmark</td><td>VIP 3000</td><td></td></tr><tr><td>Feedback regulated heating blanket</td><td>Harvard Apparatus</td><td>50-7079F</td><td></td></tr><tr><td>ECG monitor</td><td>Digicare Biomedical</td><td>LifeWindow Lite</td><td></td></tr><tr><td>EEG amplifier</td><td>A-M Systems</td><td>1800</td><td></td></tr><tr><td>EEG display monitor</td><td>Hewlett Packard</td><td>78304A</td><td></td></tr><tr><td>End tidal CO&lt;sub&gt;2&lt;/sub&gt; monitor</td><td>Respironics</td><td>Novametrix Capnoguard 1265</td><td>Optimize ventilation</td></tr><tr><td>Carbide drill burrs for drilling bone</td><td>Henry Schein</td><td>fine (0.5 mm tip) and coarse (1.25 mm tip)</td><td></td></tr><tr><td>Cement for headplate/chamber</td><td>Dentsply</td><td>675571, 675572</td><td></td></tr><tr><td>Black Powder Tempera Paint</td><td>Sargent Art Inc.</td><td>22-7185</td><td>Add to cement to improve light shielding and reduce reflections</td></tr><tr><td>Agarose - Type III-A</td><td>Sigma</td><td>A9793</td><td>For minimizing pulsations during intrinsic signal and two-photon imaging</td></tr><tr><td>Coverglass: 5 or 8 mm diameter, 0.17 mm thickness</td><td>World Precision Instruments</td><td>502040, 502041</td><td>For minimizing pulsations during imaging, the coverglass may be cut as needed</td></tr><tr><td>Brudon curettes</td><td>George Tiemann</td><td>105-715-0, 105-715-3</td><td>Cleaning skull surface</td></tr><tr><td>Bone wax</td><td>Ethicon</td><td>W31G</td><td>Quickly stop bleeding</td></tr><tr><td>Cotton Tipped Applicator</td><td>Electron Microscopy Sciences</td><td>72308-05</td><td>Clean and dry bone surface</td></tr><tr><td>Dumont #5CO Forceps</td><td>Fine Science Tools</td><td>11295-20</td><td>Grab individual layers of dura or pia</td></tr><tr><td>Vannas Spring Scissors</td><td>Fine Science Tools</td><td>15000-03</td><td>Cut dura</td></tr><tr><td>Gelfoam</td><td>Pfizer</td><td>09-0396-05</td><td>To stop bleeding on the dura</td></tr><tr><td>Absorption spears</td><td>Fine Science Tools</td><td>18105-01</td><td>Ultra-fast and lint-free wicking of CSF</td></tr><tr><td>Blackout material</td><td>Thorlabs</td><td>BK5</td><td>Shield craniotomy</td></tr><tr><td colspan="4">&lt;strong&gt;2. Dye preparation / injection&lt;/strong&gt;</td></tr><tr><td>Dimethyl Sulphoxide (DMSO)</td><td>Sigma</td><td>D2650</td><td></td></tr><tr><td>Pluronic</td><td>Sigma</td><td>P2443</td><td></td></tr><tr><td>Oregon Green 488 Bapta-1 AM</td><td>Invitrogen</td><td>O6807</td><td>Calcium indicator</td></tr><tr><td>Alexa Fluor 594</td><td>Invitrogen</td><td>A10438</td><td></td></tr><tr><td>Centrifugal filter (0.45 &amp;mu;m pore size)</td><td>Millipore</td><td>UFC30HV00</td><td>To remove impurities before injection</td></tr><tr><td>Glass pipette puller</td><td>Sutter Instruments</td><td>P97</td><td></td></tr><tr><td>Borosilicate glass filamented capillary (1.5 mm outer diameter)</td><td>World Precision Instruments</td><td>1B150F-4</td><td>Dye ejection pipette</td></tr><tr><td>Microloader</td><td>Eppendorf</td><td>5242 956 003</td><td>For loading dye into pipette</td></tr><tr><td>Micromanipulator</td><td>Sutter Instruments</td><td>MP-285</td><td>To position pipette</td></tr><tr><td>Pressure pulse controller</td><td>Parker Hannifin</td><td>PicoSpritzer III</td><td>For pressure injection of the dye</td></tr><tr><td>Single-cell electroporator</td><td>Molecular Devices</td><td>Axoporator 800A</td><td>For electroporation of the dye</td></tr><tr><td colspan="4">&lt;strong&gt;3. Intrinsic imaging&lt;/strong&gt;</td></tr><tr><td>4x Objective (0.13 NA, 17 mm WD)</td><td>Olympus</td><td>UPLFLN4X</td><td></td></tr><tr><td>Intrinsic hardware / software</td><td>Optical Imaging Inc.</td><td>Imager 3001 / VDAQ</td><td>VDAQ software is used for episodic imaging</td></tr><tr><td>CCD Camera</td><td>Adimec</td><td>Adimec-1000</td><td></td></tr><tr><td>Light source power supply</td><td>KEPCO</td><td>ATE 15-15M</td><td></td></tr><tr><td>Light source</td><td>Optical Imaging Inc.</td><td>HAL 100</td><td>Light intensity at the cortical surface is 3-5 mW</td></tr><tr><td>Green filter (for vascular image)</td><td>Optical Imaging Inc.</td><td>&amp;#955; = 546 nm (bandpass 30 nm)</td><td>For reference image of surface vasculature</td></tr><tr><td>Red filter (for intrinsic signal)</td><td>Optical Imaging Inc.</td><td>&amp;#955; = 630 nm (bandpass 30 nm)</td><td>To collect intrinsic signals</td></tr><tr><td>Heat filter</td><td>Optical Imaging Inc.</td><td>KG-1</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;4. Two-photon rig/imaging&lt;/strong&gt;</td></tr><tr><td>Two-photon microscope and software</td><td>Prairie Technologies</td><td></td><td>See Shen &lt;em&gt;et al.&lt;/em&gt; 2012 for light path, filters and laser power</td></tr><tr><td>Ti:Sapphire laser</td><td>Spectra-Physics</td><td>Mai Tai XF</td><td></td></tr><tr><td>20x (0.5 NA; 3.5 mm WD)</td><td>Olympus</td><td>UMPLFLN20X</td><td>0.5 NA objective is used only for aligning pipette over the craniotomy (not for two photon imaging)</td></tr><tr><td>20x (1.0 NA; 2.0 mm WD)</td><td>Olympus</td><td>XLUMPLFLN20X</td><td></td></tr><tr><td>40x (0.8 NA; 3.3 mm WD)</td><td>Olympus</td><td>LUMPLFLN40X/IR</td><td></td></tr><tr><td>Air table</td><td>Newport</td><td>ST-200</td><td>Isolates preparation from external vibrations</td></tr><tr><td>&lt;em&gt;xy&lt;/em&gt; stage</td><td>Mike&amp;#8217;s Machine Co. (Attleboro, MA)</td><td></td><td>Experimental subject and Sutter micromanipulator placed on &lt;em&gt;xy&lt;/em&gt; stage</td></tr><tr><td colspan="4"> &lt;table border=&quot;1&quot;&gt; &lt;tbody&gt; &lt;tr&gt; &lt;td colspan=&quot;2&quot;&gt;&lt;strong&gt;Recipes&lt;/strong&gt;&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;Artificial Cerebro-Spinal Fluid&lt;/td&gt; &lt;td&gt;NaCl (135 mM), KCl (5.4 mM), MgCl&lt;sub&gt;2&lt;/sub&gt; (1.0 mM), CaCl&lt;sub&gt;2&lt;/sub&gt; (1.8 mM), HEPES (5 mM), pH 7.4&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;Pipette Solution&lt;sup&gt;14&lt;/sup&gt;&lt;/td&gt; &lt;td&gt;NaCl (150 mM), KCl (2.5 mM), HEPES (10 mM), pH 7.4&lt;/td&gt; &lt;/tr&gt; &lt;/tbody&gt; &lt;/table&gt; </td></tr></tbody></table>

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.

To illustrate the precision of our dye labeling methods, we targeted the smallest micro-domain of any known functional map in the non-rodent neocortex. Sparsely punctuated throughout the orientation map in the primary visual cortex are singularities. These occur at points where all preferred orientations converge such that in false color maps of preferred orientation, the regions around the singularity look like "pinwheels" (Figure 2A-B). One pinwheel per craniotomy is selected for dye labeling (green circle in Figure 2C), being sure to avoid large vessels and arterioles in particular15. With bulk-loading of OGB-1 AM, tracking changes in neuronal calcium indicator fluorescence to a sequence of oriented grating visual stimuli reveals single-cell resolution functional maps around pinwheel singularities (Figure 4). With single-cell electroporation, the spatial organization of dendrites and axons from a single neuron located at a pinwheel singularity is determined in vivo (Figure 5).
<img alt="Figure 1" fo:content-width="4in" fo:src="/files/ftp_upload/50025/50025fig1highres.jpg" src="/files/ftp_upload/50025/50025fig1.jpg" /> 
Figure 1. Intrinsic signal optical imaging setup. (A) The animal is positioned on the xy stage and intrinsic signal imaging is performed via a CCD camera mounted on the two-photon microscope. The light source and computer used to control the imaging session are brought over to the setup for intrinsic imaging and moved away before the two-photon imaging phase of the experiment begins. (B) Close up view of the 4x objective and the 630 nm illumination through liquid light guides. Either two light guides attached to metal posts (left), or a single illuminating ring attached to directly the objective (right) can be used. Blackout material used to shield the objective from external sources of illumination, e.g., visual stimulus display, during optical imaging sessions, is not shown. (C) Schematic of the microscope and light path in the conventional two-photon imaging configuration. See Supplementary Fig. 20 in Shen et al. 201215 for a description of the complete two-photon light path. The retractable mirror is positioned to deflect the light from the laser through the dichroic and onto the cortex. The primary dichroic, which passes the laser beam and reflects the emitted light from the cortex to the PMT's, is moved into the light path by turning the filter turret. (D) For intrinsic imaging the mirror is retracted and the filter turret is turned to move the primary dichroic out of the way, allowing a direct light path from the cortex to the CCD camera. <a href="https://www.jove.com/files/ftp_upload/50025/50025fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
 
<img alt="Figure 2" src="/files/ftp_upload/50025/50025fig2.jpg" /> 
Figure 2. Using functional maps obtained with intrinsic signal imaging to select a micro-domain for labeling neurons with fluorescent dyes. (A) Orientation map from cat primary visual cortex acquired via intrinsic signal optical imaging. The color of each pixel represents the preferred orientation at that location. Potential orientation pinwheel singularities suitable for labeling neurons with fluorescent dyes are shown within black circles. (B) The map of cortical surface blood vessels overlaid with the orientation map. (C) Pinwheels within red circles are near large blood vessels or arterioles and so the pinwheel site within the green circle is chosen as the target for dye labeling. The dashed rectangle corresponds to the region shown in Figure 3B. Scale bar in (A) 500 &mu;m.
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/50025/50025fig3highres.jpg" src="/files/ftp_upload/50025/50025fig3.jpg" /> 
Figure 3. Optimizing the pipette placement for loading neurons with fluorescent dyes. (A) Schematic side-view of the recording chamber shows that the dye loading site to be targeted in visual cortex (red-and-white bull's eye) is ~200 &mu;m below the cortical surface. The xy coordinate of the target corresponds to the location of an orientation pinwheel singularity determined from intrinsic signal imaging. The pipette will enter the cortex at a 30&deg; angle. If the cortical surface is parallel to the xy stage surface, the pipette tip will move laterally approximately 350 &mu;m before reaching the pinwheel singularity. (B) Top view of recording chamber shows the cortical surface vasculature and the pipette entry position (red star) that is 350 &mu;m from the pinwheel singularity. (C) The xy stage position is adjusted such that the pipette entry position is centered under the 20x or 40x objective (3.3-3.5 mm WD). (D) The objective is raised ~2 mm. (E) The pipette is brought over the craniotomy and its tip is positioned directly above the entry position on the cortical surface using green epi-fluorescent light. (F) The pipette and objective are lowered concurrently using bright-field illumination until the cortical surface blood vessels come into focus. (G) The objective and ACSF are removed and a drop of 3% agarose is applied over the craniotomy. (H) Two-photon visualization with a 40x (0.8 NA, 3.3 mm WD) objective is used to follow the pipette down to the chosen cortical depth and inject the dye. After confirming that neurons are successfully labeled with fluorescent dye, typically, a higher NA 20x objective (1.0 NA, 2.0 mm WD) is used to collect two-photon t- and z-series data. Schematics shown in A, C-H are not drawn to scale.
<img alt="Figure 4" src="/files/ftp_upload/50025/50025fig4.jpg" /> 
Figure 4. Correspondence of orientation preference in two-photon and intrinsic signal imaging functional maps. (A) Two-photon imaged area showing cell bodies labeled with OGB-1 AM in visual cortex layer 2/3. (B) Orientation preference of individual neuronal cell bodies from the same site shown in (A). Around the pinwheel singularity that was centered in the imaged region is an organized map of preferred stimulus orientation. (C) The orientation map obtained with intrinsic signal imaging. The black square corresponds to the region shown in (B). These data were obtained from the same experiment shown in Figures 2A-C and 3B. Scale bar in (A-B) 100 &mu;m and (C) 500 &mu;m.
<img alt="Figure 5" src="/files/ftp_upload/50025/50025fig5.jpg" /> 
Figure 5. In vivo morphology of a single neuron at an orientation pinwheel singularity. (A) The dendrites and cell body of a single neuron overlaid with the orientation map from the same visual cortical site. The neuron was electroporated with Alexa Fluor 594 under two-photon guidance, a z-stack was collected in vivo, and dendritic processes were traced offline using Neurolucida software. The orientation map was obtained using intrinsic signal imaging before single-cell electroporation. (B) Average intensity 10-&mu;m-z-projection from cortical layer 1 shows apical dendrites. (C-D) Average intensity 10-&mu;m-z-projections from cortical layer 2/3 show basal dendrites and axons. Red arrows show spines along dendrites and white arrows point to individual axons. Scale bars in (A-D) 100 &mu;m.

Watch this Scientific Journal Video about Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging at JoVE.com

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

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, ...

targeted-labeling-neurons-specific-functional-micro-domain-neocortex

Nanyang Technological University

Research

JoVE Journal

Neuroscience

13.6K Views.  Medical University of South Carolina. 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.

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

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

Biology

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 ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures in behaving animals remains a technical challenge, especially if the structures must be targeted stereotaxically. This protocol describes convenient surgical and electrophysiological techniques that maintain the animal&#8217;s head in the stereotaxic plane and unambiguously isolate the spiking activity of single neurons. The protocol combines head restraint of alert rodents, juxtacellular monitoring with micropipette electrodes, and iontophoretic dye injection to identify the neuron location in post-hoc histology. While each of these techniques is in itself well-established, the protocol focuses on the specifics of their combined use in a single experiment. These neurophysiological and neuroanatomical techniques are combined with behavioral monitoring. In the present example, the combined techniques are used to determine how self-generated vibrissa movements are encoded in the activity of neurons within the somatosensory thalamus. More generally, it is straightforward to adapt this protocol to monitor neuronal activity in conjunction with a variety of behavioral tasks in rats, mice, and other animals. Critically, the combination of these methods allows the experimenter to directly relate anatomically-identified neurophysiological signals to behavior.

This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical properties, synaptic connections, and in vivo functions, but their basic functional and anatomical organization from cellular to network scale is poorly understood. Here we describe a method for the three-dimensional imaging of fluorescently-labeled neurons across large areas of the brain for the investigation of the cortical cellular organization. Specific types of neurons are labeled by the injection of fluorescent retrograde neuronal tracers or expression of fluorescent proteins in transgenic mice. Block brain samples, e.g., a hemisphere, are prepared after fixation, made transparent with tissue clearing methods, and subjected to fluorescent immunolabeling of the specific cell types. Large areas are scanned using confocal or two-photon microscopes equipped with large working distance objectives and motorized stages. This method can resolve the periodic organization of the cell type-specific microcolumn functional modules in the mouse neocortex. The procedure can be useful for the study of three-dimensional cellular architecture in the diverse brain areas and other complex tissues.

Here we describe a procedure for tissue clearing, fluorescent labeling, and large-scale imaging of mouse brain tissue which, thereby, enables visualization of the three-dimensional organization of cell types in the neocortex.

The mammalian neocortex is composed of many types of excitatory and inhibitory neurons, each with specific electrophysiological and biochemical ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

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.

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

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Large-scale Three-dimensional Imaging of Cellular Organization in the Mouse Neocortex

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

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