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 appl...

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalogue Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>1. Life support/experiment prep</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Webster Vet</td>
			<td>NDC 57319-474-05</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Midmark</td>
			<td>VIP 3000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Feedback regulated heating blanket</td>
			<td>Harvard Apparatus</td>
			<td>50-7079F</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>ECG monitor</td>
			<td>Digicare Biomedical</td>
			<td>LifeWindow Lite</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>EEG amplifier</td>
			<td>A-M Systems</td>
			<td>1800</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>EEG display monitor</td>
			<td>Hewlett Packard</td>
			<td>78304A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>End tidal CO2 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>&nbsp;</td>
		</tr>
		<tr>
			<td>Cement for headplate/chamber</td>
			<td>Dentsply</td>
			<td>675571, 675572</td>
			<td>&nbsp;</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>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>2. Dye preparation / injection</td>
		</tr>
		<tr>
			<td>Dimethyl Sulphoxide (DMSO)</td>
			<td>Sigma</td>
			<td>D2650</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Pluronic</td>
			<td>Sigma</td>
			<td>P2443</td>
			<td>&nbsp;</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>&nbsp;</td>
		</tr>
		<tr>
			<td>Centrifugal filter (0.45 &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>&nbsp;</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>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>3. Intrinsic imaging</td>
		</tr>
		<tr>
			<td>4x Objective (0.13 NA, 17 mm WD)</td>
			<td>Olympus</td>
			<td>UPLFLN4X</td>
			<td>&nbsp;</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>&nbsp;</td>
		</tr>
		<tr>
			<td>Light source power supply</td>
			<td>KEPCO</td>
			<td>ATE 15-15M</td>
			<td>&nbsp;</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>&lambda; = 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>&lambda; = 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>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>4. Two-photon rig/imaging</td>
		</tr>
		<tr>
			<td>Two-photon microscope and software</td>
			<td>Prairie Technologies</td>
			<td>&nbsp;</td>
			<td>See Shen et al. 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>&nbsp;</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>&nbsp;</td>
		</tr>
		<tr>
			<td>40x (0.8 NA; 3.3 mm WD)</td>
			<td>Olympus</td>
			<td>LUMPLFLN40X/IR</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Air table</td>
			<td>Newport</td>
			<td>ST-200</td>
			<td>Isolates preparation from external vibrations</td>
		</tr>
		<tr>
			<td>xy stage</td>
			<td>Mike's Machine Co. (Attleboro, MA)</td>
			<td>&nbsp;</td>
			<td>Experimental subject and Sutter micromanipulator placed on xy stage</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>
			<table border="1">
				<tbody>
					<tr>
						<td colspan="2">Recipes</td>
					</tr>
					<tr>
						<td>Artificial Cerebro-Spinal Fluid</td>
						<td>NaCl (135 mM), KCl (5.4 mM), MgCl2 (1.0 mM), CaCl2 (1.8 mM), HEPES (5 mM), pH 7.4</td>
					</tr>
					<tr>
						<td>Pipette Solution14</td>
						<td>NaCl (150 mM), KCl (2.5 mM), HEPES (10 mM), pH 7.4</td>
					</tr>
				</tbody>
			</table>
			</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 ci...

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

Research

JoVE Journal

Neuroscience

13.6K Aufrufe.  Medical University of South Carolina. Es wird ein Verfahren zur Markierung von Neuronen mit Fluoreszenzfarbstoffen in vorbestimmten funktionellen Mikrodomänen des Neokortex beschrieben. Zuerst wird intrinsische Signal Abbildungsoptik verwendet werden, um eine funktionelle Karte zu erhalten. Dann Zweiphotonenmikroskopie wird Etiketts und Bild Neuronen innerhalb eines Mikro-Domäne von der Karte verwendet.

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

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

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 ...

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 ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

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+ ...

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 ...

Advanced Diffusion Imaging in The Hippocampus of Rats with Mild Traumatic Brain Injury

Mild traumatic brain injury (mTBI) is the most common type of acquired brain injury. Since patients with traumatic brain injury show a tremendous variability and heterogeneity (age, gender, type of trauma, other possible pathologies, etc.), animal models play a key role in unraveling factors that are limitations in clinical research. They provide a standardized and controlled setting to investigate the biological mechanisms of injury and repair following TBI. However, not all animal models mimic the diffuse and subtle nature of mTBI effectively. For example, the commonly used controlled cortical impact (CCI) and lateral fluid percussion injury (LFPI) models make use of a craniotomy to expose the brain and induce widespread focal trauma, which are not commonly seen in mTBI. Therefore, these experimental models are not valid to mimic mTBI. Thus, an appropriate model should be used to investigate mTBI. The Marmarou weight drop model for rats induces similar microstructural alterations and cognitive impairments as seen in patients who sustain mild trauma; therefore, this model was selected for this protocol. Conventional computed tomography and magnetic resonance imaging (MRI) scans commonly show no damage following a mild injury, because mTBI induces often only subtle and diffuse injuries. With diffusion weighted MRI, it is possible to investigate microstructural properties of brain tissue, which can provide more insight into the microscopic alterations following mild trauma. Therefore, the goal of this study is to obtain quantitative information of a selected region-of-interest (i.e., hippocampus) to follow up disease progression after obtaining a mild and diffuse brain injury.

The overall goal of this procedure is to obtain quantitative microstructural information of the hippocampus in a rat with mild traumatic brain injury. This is done using an advanced diffusion-weighted magnetic resonance imaging protocol and region-of-interest based analysis of parametric diffusion maps.

Mild traumatic brain injury (mTBI) is the most common type of acquired brain injury. Since patients with traumatic brain injury show a tremendous ...

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...