This study was supported by a grant of the German Science Foundation (DFG, Ro2327/6-1) to C.R.R. The authors wish to thank S. Durry and C. Roderigo for expert technical assistance, and M. D&#252;bbert (Electronics Laboratory, Zoological Institute, University of Cologne, Germany) for help in implementing the Pockels cell controller and the HF switch.

This study was carried out in strict accordance with the institutional guidelines of the Heinrich Heine University D&#252;sseldorf, Germany, as well as the European Community Council Directive (86/609/EEC). All experiments were communicated to and approved by the Animal Welfare Office at the Animal Care and Use Facility of the Heinrich Heine University D&#252;sseldorf, Germany (institutional act number: O52/05). In accordance with the German Animal Welfare Act (Tierschutzgesetz, Articles 4 and 7), no formal additional approval for the postmortem removal of brain tissue was necessary.
For generation of acute slices, mice were anaesthetized with CO2 and quickly decapitated (following the recommendation of the European Commission published in: Euthanasia of experimental animals, Luxembourg: Office for Official Publications of the European Communities, 1997; ISBN 92&#8211;827-9694-9).
1. Preparation of Solutions
<ol>
	<li>Prepare artificial cerebrospinal fluid (ACSF) for dissection containing (in mM): 125 NaCl, 2.5 KCl, 0.5 CaCl2, 6 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, and 20 glucose, bubbled with 95% O2 and 5% CO2, resulting in a pH of 7.4.</li>
	<li>Prepare ACSF for experiments containing (in mM): 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgCl2, 1.25 NaH2PO4, 26 NaHCO3, and 20 glucose, bubbled with 95% O2 and 5% CO2, resulting in a pH of 7.4.</li>
	<li>Prepare intracellular solution (ICS) containing (in mM): 150 KMeSO3, 12.5 hydroxy ethyl piperazine ethane sulfonic acid (HEPES), 40 KCl, 5 NaCl, 1.25 ethylene glycol tetraacetic acid (EGTA), 5 Mg-ATP, 0.5 Na-GTP. Adjust pH to 7.3. Store aliquots of 1 ml at -20 &#176;C.</li>
	<li>Dilute sodium-binding benzofuran isophthalate (SBFI)-salt in double-distilled water and prepare aliquots of 5 &#181;l of a 10 mM stock solution.</li>
	<li>Thaw ICS and add SBFI stock solution at a final concentration of 1 mM. Vortex and micro-filtrate (0.22 &#181;m). Keep at 4 &#176;C until used in experiment. Do not refreeze and only use for one day.</li>
	<li>Prepare MNI glutamate stock solution of 50 mM by dissolving the compound in double-distilled water. Dilute MNI glutamate stock solution to a final concentration of 5 mM in normal ACSF. Keep at 4 &#176;C until used in experiment. Do not refreeze and only use for one day. Store aliquots, which are not immediately used in the experiment, at -20 &#176;C.</li>
</ol>
2. Dissection of Tissue
NOTE: The preparation of acute hippocampal slices of the rodent brain was described in detail earlier13,14. In brief, the following protocol was employed in the present study.
<ol>
	<li>After decapitation, rapidly remove the brain from the skull.</li>
	<li>Immediately place the brain in a petri dish with ice-cold dissection ACSF and dissect a hemisphere by performing a sagittal cut along the midline.</li>
	<li>Perform a second cut in the desired orientation as a blocking surface and attach this to the cutting stage of a vibratome with superglue.</li>
	<li>Take cutting chamber and cooling element (both kept at -20 &#176;C) of the vibratome out of the freezer and place cutting stage with brain section in the chamber. Then put cooling element in the chamber and submerge tissue in ice-cold dissection ACSF. To stabilize the preparation, one may want to counter the tissue block with agar gel.</li>
	<li>Cut 250 &#181;m thick, parasagittal slices of the hippocampus with the vibratome. Ensure that all ACS fluids are bubbled at all times.</li>
	<li>After slicing, keep tissue on a mesh in a beaker with the normal ACSF and incubate at 34 &#176;C for 30 min. Then keep at room temperature.</li>
</ol>
3. Preparation of Hardware
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/52038/52038fig1highres.jpg" width="500" /> 
Figure 1. Scheme showing light paths and experimental control of the rig consisting of multi-photon imaging, laser-scanning UV flash photolysis, and electrophysiology. The multi-photon beam (red) is produced by a pulsed, tunable infrared (IR) laser (TiSa). It passes a mechanical shutter, a Pockels&#8217; cell, and the IR detector (detection of beam intensity), all of which enable control of the laser power and its administration duration. The flip-in/flip-out optional laser diode is employed for basic alignment of the laser beam. The beam expander may be used in combination with objectives with an extremely wide back-focal plane. The remote-controlled ND filter wheel may be used in addition or instead of the Pockels&#8217; cell to control laser beam power. After passing the scan head, the pulsed IR-light is guided to the specimen. Emitted fluorescence light (light green) is collected either by the external or the internal photomultiplier detectors (PMTs). The external detectors are fully synchronized with the internal PMTs via a high frequency switch (PMT controller). The uncaging beam (light blue) is produced by a UV solid-state laser (DPSS UV-Laser). It is then directed to the galvo-driven scanning unit at the top-back of the epi-fluorescence condenser by a light guide. The precise positioning (chromatic aberration between imaging and uncaging beam) of the uncaging spot or area is enabled by the image export from the imaging software. The timing management for synchronizing the imaging, the electrophysiology, and the flash photolysis is electronically controlled. The transillumination detector (TL-PMT) is of need for documentation of the positioning of the pipettes within the tissue. The control units and software of the imaging system, which has been modified, is used to control and synchronize all other devices needed to run the system. System components labeled as &#8220;custom&#8221; were designed and build/adapted by the authors. Some components were adapted to meet the requirements of the custom-build multi-photon system and are labeled as &#8220;modified&#8221;. <a href="https://www.jove.com/files/ftp_upload/52038/52038fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Switch on components of the multi-photon system. Test and adjust infrared laser beam alignment.
	<ol>
		<li>Control beam positioning by flipping in the centering prism instead of an objective. If the beam is dislocated, re-adjust it by the mirrors in the periscope. Note that the centering plane of the prism must be at the same level as the back focal plane of the objective while imaging.</li>
		<li>Switch on the spectrometer and check for the multi-photon characteristics of the beam.</li>
		<li>Set parameters of the imaging software to the following values: Choose a frame size of 512 x 512 pixels for overview images of the cell (smaller clip boxes with a zoom factor of 2.5 for high frame rate and high spatial resolution, respectively). Set imaging speed to fast mode and scan mode to XYT. Z-stepping should be 1 &#181;m for overview stacks and 0.2 &#181;m for detailed stacks to match spatial sampling requirements for deconvolution performed later.</li>
	</ol>
	</li>
	<li>Adjust multi-photon laser beam (790 nm) intensity by altering settings of the Pockels&#8217; cell (final power under the objective: &#8776; 14 - 16 &#181;W) and the photo-multipliers (PMTs).</li>
	<li>Switch on and calibrate uncaging system by placing a fluorescent sample slide under the objective lens.
	<ol>
		<li>Using the binoculars, adjust the focus of the UV optics and spatially limit the UV laser spot to a size of 2 &#181;m in diameter at maximum by employing the focusing unit at the UGA scan head.</li>
		<li>Start the calibration routine of the uncaging unit control software.
		<ol>
			<li>Set the UV laser to several points within its scan range, while the fluorescence is captured with a CCD camera. By clicking on the UV laser spot at each point, adjust the positioning of the galvanic scan mirrors to correspond to a certain coordinate in the software.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/52038/52038fig2highres.jpg" width="500" /> 
Figure 2. Detailed scheme illustrating the characteristics ofexcitation, emission and uncaging beam paths. The IR excitation beam (red) passes the beam combining dichroic mirror at the level to the filer turret in the epi-fluorescence condenser of the microscope and the objective to reach the specimen. The uncaging beam (blue) is delivered via a quartz optical light fiber to the collimation and beam focusing unit and then subsequently guided to the scanning mirrors in the scanning unit, passes a dichroic mirror to the beam combining dichroic mirror. Here it is combined with the excitation scanning beam. The emitted light from the specimen follows the path of the excitation scanning beam path in the opposite orientation towards the imaging scan head. The camera is used for visual control of the patch pipette and for positioning of the uncaging beam in combination with the confocal laser scanning microscope (not shown). The green light path represents an optional epi-fluorescence illumination which may be of use with other applications.
<ol start="3">
	<li>
	<ol start="2">
		<li>
		<ol start="2">
			<li>Use the flash photolysis system in the spot mode to gain focal applications. Ensure that the focal adjustment of the uncaging beam (average power underneath the objective: 0.55 mW) results in a spot size of 1.5 &#181;m in diameter (xy extension) as revealed at a fluorescent slide positioned at a z-level that corresponds to that of a tissue slice (not shown).</li>
		</ol>
		</li>
		<li>The accurate positioning of the uncaging spot is achieved by the import of an image of the imaging system via a network connection between uncaging- and imaging-computers.
		<ol>
			<li>Using a screen grabber software, continuously read out the frames of the imaging software (instead of the camera feed) and adjust the imaging and uncaging frames congruently. Export every 10th frame from the imaging software as a reference image into the flash unit to ensure proper adjustment during the entire experiment.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" fo:content-width="6.5in" src="/files/ftp_upload/52038/52038fig3highres.jpg" width="600" /> 
Figure 3. Adjustment of the uncaging spot: To precisely position the uncaging spot, a screen shot of the imaging software (yellow frame) is imported into the uncaging software (green frame). The left image within the yellow frame represents a Hi-Lo coded image (blue: black pixels, red: saturated pixels) of the entire CA1 pyramidal cell. The image is overlaid by the calibration grid of the uncaging software (green &#8220;X&#8221;s). To the right, a magnified part of the cell is shown with the overlaid uncaging spot (red cross). The yellow spot is the automatically overlaid image of the uncaging beam. <a href="https://www.jove.com/files/ftp_upload/52038/52038fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Turn on micromanipulators, electrophysiology components, and pressure application device for delivery of the caged compound to the target region.</li>
	<li>Install a focal pressure application device for local perfusion of caged compounds. This will reduce costs enormously as compared to bath perfusion of these substances. Adjust the holding pressure to the given atmospheric pressure to prevent a drag of ACSF into or a leakage of caged substances from the application pipette. 
	NOTE: The pressure application device hosts a highly precise and ultrafast micro-valve in the pipette holder. This enables to employ minimal application pressures and therefore reduces movement artifacts to a minimum during the local perfusion with the caged compound.</li>
	<li>Pull pipettes for whole-cell patch-clamp and local perfusion using fire-polished borosilicate glass capillaries. Pipettes should have a tip diameter of ~1 &#181;m and a resistance of R &#8776; 3 M&#8486; (values determined with K-MeSO3-based ICS).</li>
	<li>Place slice in the experimental bath and affix it with a grid (platinum frame 250 &#181;M thick, spanned with surgical filaments, o.d. 40 &#181;m) (Figure 4A, B). Use an inverted, tip-broken, and fire-polished Pasteur pipette (suction ball at the side of the broken tip) for slice transfer. Avoid bending and any other harsh handling of the slice.</li>
</ol>
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/52038/52038fig4highres.jpg" width="500" /> 
Figure 4. Components of the experimental bath and positioning of the bath at the microscope stage. (A) The experimental bath consists of the bath chamber itself (blue square), which is surrounded by a magnetic metal ring. The tubing ensures continuous perfusion of the bath with saline (ACSF). The grid to hold down and fix the slice is put into the bath chamber. (B) Acute slice preparation positioned within the bath chamber. The slice is fixed by the threads of the grid. (C) Positioning and geometry of the experimental bath at the microscope stage during the experiments.
<ol start="8">
	<li>Place bath at the microscope stage and permanently perfuse slice with ACSF.</li>
</ol>
4. Whole-cell Patch-clamping
<ol>
	<li>Load patch pipette with ICS containing SBFI and load local perfusion pipette with caged compound. Attach pipettes to corresponding micromanipulators. Place reference electrode in bath. Make sure that you are grounded permanently to avoid damage to the head stage (c.f. Figure 4C).</li>
	<li>Lower both pipettes into bath and place them above the hippocampal CA1 region. Apply gentle pressure to patch pipette (+40 mbar) to avoid dilution of the ICS with ACSF.</li>
	<li>Compensate the offset potential of the patch pipette using the electrophysiology software.</li>
	<li>Approach a CA1 pyramidal cell with patch pipette employing IR-DIC video microscopy. Apply gentle suction until a Giga-seal is obtained. Choose a cell the soma of which is located 30 -70 &#181;m below the surface of the slice to ensure intact cell morphology on the one hand side and low scattering and attenuation of the uncaging beam on the other hand side.</li>
	<li>Compensate for fast capacity. Break membrane and open cell to gain whole-cell configuration.</li>
	<li>Compensate for slow capacity and series resistance.</li>
	<li>Allow the cell to be dialyzed with the SBFI-ICS for at least 30 min before starting the imaging experiments.</li>
</ol>
5. Multi-photon imaging and stimulation
<img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/52038/52038fig5highres.jpg" width="600" /> 
Figure 5. Experimental protocol. To initialize an experiment, a trigger pulse (indicated by red sign (&#7464;)and the red line) is set to simultaneously start the imaging (blue), the electrophysiology (green), and the administration of the caged compound (yellow) at time point 0 sec. During this first period, the imaging beam should be entirely dimmed (light blue). After 4.5 sec (dashed line), the local perfusion of the caged compound is terminated and the imaging beam should be set to its working intensity for data acquisition (blue). 1.5 sec afterwards (time point 6 sec), a second trigger pulse is given (indicated by red sign (&#7464;) and the red line), which initializes the UV flash unit (duration of the flash: 300 msec; indicated by the yellow flash) and sets markers in the electrophysiology and the imaging protocol.
<ol>
	<li>Add tetrodotoxin (TTX, 500 nM) to ACSF to prevent activation of voltage-gated sodium channels and generation of action potentials.</li>
	<li>Visualize cellular morphology using multi-photon excitation and resulting SBFI fluorescence and choose a spiny dendrite for experiment. Zoom in for images at a higher resolution and place a clip box around the dendrite.</li>
	<li>Fill a standard patch pipette with 10 &#181;l ACSF containing caged glutamate. Connect the pipette to the pressure application system and attach it to the micromanipulator.</li>
	<li>Place pipette with caged compound near the dendrite (~30 &#181;m). Position the pipette to allow efficient local perfusion of the dendrite of choice. Adjust the uncaging laser: position the uncaging spot close (~1 - 2 &#181;m) to the structure of interest.</li>
	<li>Set regions of interest on the chosen dendrite and adjacent spines using imaging software.</li>
	<li>Approach the region of interest and focally inject the caged compound for several sec with low pressure (&#60;3 PSI). Start patch clamp and fluorescence recordings (at 790 nm multi-photon excitation) via trigger signal. 
	NOTE: During the local perfusion of the caged compound, the excitation beam is completely dimmed to prevent bleaching.</li>
	<li>Stop local perfusion of the caged compound. Increase intensity of the two-photon laser to enable efficient excitation of SBFI and apply a UV flash to initialize uncaging (uncaging duration 300 msec).</li>
	<li>Monitor changes in SBFI fluorescence. Stop recording after SBFI fluorescence has recovered back to baseline.</li>
</ol>
6. Pharmacology
<ol>
	<li>To test the involvement of ionotropic glutamate receptors in the generation of the currents and/or sodium signals evoked by UV flash photolysis of caged glutamate, employ receptor blockers.
	<ol>
		<li>Switch to ACSF containing the AMPA-receptor blocker cyano-nitroquinoxaline-dione (CNQX, 10 &#181;M) and the NMDA-receptor blocker amino-phosphonopentanoate (APV, 50 &#181;M) in addition to TTX (see above) and perfuse slice for at least 10 min.</li>
		<li>Repeat stimulation procedure (see 5.4 and 5.5.)</li>
	</ol>
	</li>
	<li>Reversibility of the inhibitors&#8217; effects
	<ol>
		<li>Switch back to normal ACSF containing only TTX.</li>
		<li>Perfuse slice for 20 min.</li>
		<li>Repeat stimulation procedure (see 5.4 and 5.5.).</li>
	</ol>
	</li>
</ol>
7. Morphology
<ol>
	<li>Record a XYZ-stack of the clip box-region set for the performed measurements. Ensure that this stack is oversampled spatially (at least 0.2 &#181;m per pixel) to enable optimal image deconvolution and to increase image quality and resolution.</li>
	<li>Record a XYZ-stack of the entire cell to assess cell morphology.</li>
	<li>Run deconvolution algorithm.</li>
</ol>

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Neurosci. Meth. 180, 9-21 (2009).</li><li>Nikolenko, V., Peterka, D., Araya, R., Woodruff, A., Yuste, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+light+modulator+microscopy.">Spatial light modulator microscopy.</a> Cold Spring Harbor protocols. , 1132-1141 (2013).</li></ol>

The authors declare no competing interests. The authors received financial support enabling Open Access Publication by Rapp OptoElectronic (Wedel, Germany), which produces an instrument used in the video article. The company was neither involved in the experiments nor in the data handling nor in the manuscript writing.

The present study shows that SBFI is well suited for two-photon imaging of intracellular sodium transients in small cellular compartments. It has to be kept in mind, however, that the quantum efficiency of SBFI is rather low15 and relative changes in the sodium concentration are quite small with physiological activity. Thus, high-resolution measurement of Na+ transients in fine processes is a relatively tedious task, and binning or averaging of several trials may be necessary to obtain satisfactory signals. In addition, the kinetics of sodium transients are surprisingly slow, making it obligatory to record for extended time periods. This observation corresponds to those in earlier studies, where monoexponential decay of sodium transients in neuronal dendrites and in astrocytes was characterized by large decay time constants in the range of 10 sec at room temperature1,2,6. Sodium transients thus seem to exert a much slower time course and much larger decay time constants as compared to calcium transients16.
Set aside these drawbacks, sodium imaging proves to be a valuable tool for the investigation of physiological properties of neuronal subdomains. For example, sodium imaging can serve to monitor excitatory synaptic activity at or close to active synapses. Because sodium is essentially not buffered8,17, activity-induced sodium transients are linearly related to a wide range of synaptic glutamate release or exogenously applied glutamate. They hence represent direct and unbiased indicators of neuronal glutamatergic activity. Moreover, sodium indicator dyes exhibit high Kd&#8217;s (SBFI&#8217;s Kd is in the range of 25 mM1). Even at the relatively high intracellular dye concentrations used to achieve sufficient brightness (usually 0.5 - 1 mM), the high Kd&#8217;s imply that the dyes themselves do not act as buffers for sodium. Consequently, they do not distort the amplitude nor time course of sodium transients, which is always a concern when calcium-sensitive dyes are introduced into the cells18. It can thus be assumed that the detected signals represent a good measure of the &#8220;real&#8221; changes in intracellular sodium. Their slow time course then implies that the velocity for intracellular diffusion for sodium in neurons is much smaller than previously assumed19.
A critical requirement for the successful execution of experiments addressing the properties of excitatory synaptic transmission and sodium influx pathways into spines and dendrites is the induction of a fast and highly localized activation of postsynaptic structures. This can be obtained by fast and local application of glutamate or glutamate agonists in the direct vicinity of a spine or a dendrite by the flash photolysis of caged compounds. Flash photolysis does not mechanically damage the tissue, is free of movement artifacts and is well established for the investigation of related questions (e.g. local calcium signaling). The diameter of the uncaging spot was 1.5 &#181;m in the xy-plane. Uncaging close to a spiny dendrite induced sodium signals in both spines and the parent dendrite. Whereas the signals tended to be largest in spines closest to the uncaging spot, the amplitudes in the parent dendrite and spines further away were still rather similar to those in direct vicinity of the uncaging spot. Uncaging was performed for 300 msec during which inward currents increased in amplitude. Uncaged glutamate might have diffused in the tissue during the same time period, activating ionotropic glutamate receptors and sodium influx on dendritic regions and spines further away from the uncaging spot.
An intrinsic problem that occurs when using a UV laser for photolysis of caged compounds administered through the cell bathing solution is &#8216;inner filtering&#8217;. Because of the high absorption rate of the cage, UV light is attenuated strongly along the way from the objective to the stimulation site20,21. A feasible way to avoid this is local perfusion with the cage that is restricted only to the stimulation site, which furthermore decreases the amount of caged compound needed to a minimum. As compared to UV laser-scanning-mediated uncaging, near-infra-red two-photon uncaging22 is spatially more precise, mainly because the accuracy of the stimulation is increased in the z-axis. On the other hand, due to the small cross-section of commonly used cages for longer wavelengths as employed with two-photon excitation, either a high concentration of the caged compound or very high, potentially phototoxic light intensities are needed. Also, two-photon uncaging necessitates a much higher investment in the equipment; among others, an additional IR laser plus the required optical components, are needed.
Both SBFI and MNI-glutamate are excitable in the identical wavelength range. This may lead to an unintended interdependency of UV uncaging laser and SBFI or IR imaging laser and MNI-glutamate, resulting in bleaching of SBFI by the uncaging flash and /or a continuous uncaging of glutamate by the IR beam. However, as shown in Figure 6C, neither a UV flash by itself nor the multi photon imaging caused such reciprocal effects under our experimental conditions.
UV-flash systems similar to the one described here are used routinely in many other laboratories and can relatively easily be incorporated into any existing multi-photon imaging microscope. It offers the advantage that the laser beam for photolysis can be positioned freely in the field of view. In addition, the system enables a fast and automated repositioning of the laser beam and the release volume, respectively, in the field of view during an experiment. Our modification simplifies and improves the accurate positioning of the uncaging spot, so that the frames of the imaging software can serve to adjust imaging and uncaging frames congruently.
Taken together, whole-cell patch-clamp and multi-photon sodium imaging in dendrites and spines of central neurons, combined with a modified procedure for UV-light-induced uncaging of glutamate, allows reliable and focal activation of glutamate receptors in the tissue. It can thus serve to analyze the properties of excitatory synaptic transmission and of postsynaptic sodium signals in neurons in the intact tissue.

Recent improvements in light microscopic techniques such as multi-photon microscopy have enabled the study of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, these techniques are now widely used to analyze activity-related electrical signals on neurons as well as concomitant calcium signals in small subcellular compartments, namely in fine dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces sodium signals in dendrites and spines, the properties of which are largely unexplored. Such signals can be analyzed by two-photon imaging of intracellular sodium ([Na+]i) which enables the on-line measurement of [Na+]i transients for prolonged periods without significant dye bleaching or photo-damage1,2.
For imaging of [Na+]i, only a few chemical indicator dyes are available, e.g. CoroNa Green or Asante Natrium Green3,4. The most commonly used fluorescent probe for Na+ imaging is sodium-binding benzofuran isophthalate (SBFI). It is a ratiometric, UV-excited dye similar to the well-known calcium-sensitive dye fura-2 and has been employed for conventional Na+ imaging in many cell types (e.g.5,6). There are different possibilities of exciting the dye and collecting its fluorescence. If high temporal resolution (i.e. a high imaging frame rate) is required in combination with spatial information, SBFI can be excited with a xenon arc lamp or a high power light-emitting diode (LED) device and its emission detected with a high speed charge-coupled device (CCD) camera7,8. For maximal spatial resolution deep in the tissue, multi-photon laser scanning microscopy is the method of choice9. The relatively low quantum efficiency of SBFI necessitates relatively high dye concentrations (0.5 - 2 mM), and direct loading of the membrane-impermeable form of SBFI via a sharp microelectrode10 or patch pipette1.
Using SBFI, earlier work performed in acute tissue slices of the rodent hippocampus demonstrated activity-related sodium transients in dendrites and spines of CA1 pyramidal neurons which are mainly caused by influx of sodium through ionotropic NMDA receptors1,2. For the study of the properties of such local sodium signals in more detail, specific activation of postsynaptic receptors by application of receptor agonists is a well-suited method of choice. To mimic presynaptic activity and transmitter release, application should be relatively brief and focused to enable local stimulation. This, however, proves to be quite challenging in the intact tissue. Local pressure application of receptor agonists using a fine-tipped pipette enables very focal application, but hosts the potential risk for producing movement of the structure of interest (e.g. such as a dendrite or dendritic spines), and thus hinders high-resolution imaging. The suitability of iontophoresis of neuro-active substances depends on their electrical properties and high current amplitudes may produce cellular artifacts as well.
One way to circumvent these obstacles is the employment of photo-activated compounds and their flash photolysis. Basically, two different principles are used for photo-activation of caged substances: I) Wide-field flash photolysis11 and II) focal uncaging employing scanning modules in combination with lasers12. While wide-field flash photolysis is used to activate larger regions of interest, e.g. an entire cell, focal uncaging is employed to specifically stimulate small cellular compartments. In the present study we demonstrate a procedure for whole-cell patch-clamp and multi-photon sodium imaging in dendrites and spines of central neurons, combined with a modified procedure for UV-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue.

<table border="0" cellpadding="0" cellspacing="0" width="1138">
	<tbody>
		<tr height="33">
			<td height="33" style="height:33px;width:239px;">Hadrware/Software Component</td>
			<td style="width:231px;">Manufacturer</td>
			<td style="width:183px;">Type</td>
			<td style="width:487px;">Comment</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Borosilicate-glass capillaries</td>
			<td>Hilgenberg</td>
			<td>1405059</td>
			<td>75 mm x 2 mm, wall thickness 0.3 mm</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Camera</td>
			<td>Jenoptik</td>
			<td>ProgRes MF</td>
			<td>IR-DIC compatible camera</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Deconvolution software</td>
			<td>SVI</td>
			<td>Huygens Pofessional X11</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Fiber optic spectrometer</td>
			<td>Ocean Optics</td>
			<td>USB 4000</td>
			<td>Miniature fiber optic spectrometer</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Filter tubes</td>
			<td>VWR</td>
			<td>cenrifugal filter, 0.2 &#181;m</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Imaging software</td>
			<td>Olympus Europe</td>
			<td>Flowview Ver 5.0c, Tiempo</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">IR beam power controller</td>
			<td>Custom built</td>
			<td></td>
			<td>Laser beam intensity control + fast switch to zero on flyback via pockels cell</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">IR laser</td>
			<td>SpectraPhysics</td>
			<td>Mai-Tai VF-N1S</td>
			<td>fs-pulsed Ti:Sa tunable laser (710-990 nm); CAUTION</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">IR power monitor</td>
			<td>Custom built</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Micromanipulator</td>
			<td>Burleigh</td>
			<td>PCS 5000</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Microscope/Imaging System</td>
			<td>Olympus Europe</td>
			<td>BX51WI/FV300</td>
			<td>Both, the microscope and the imaging systems were strongly modified</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">ND filter wheel</td>
			<td>Newport</td>
			<td>50Q04AV.2</td>
			<td>UV fused silica, optical density 0.05-2.0</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Objective</td>
			<td>Nikon Instruments Europe</td>
			<td>NIR Apo 60x/1.0w</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Patch clamp amplifier</td>
			<td>HEKA</td>
			<td>EPC-10</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Pipette puller</td>
			<td>Narishige</td>
			<td>Model PP-830</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Pneumatic drug ejection system</td>
			<td>NPI Electronic</td>
			<td>PDES-DXH</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Pockels cell</td>
			<td>Conoptics</td>
			<td>350-80 LA-BK</td>
			<td>EOM, used as fast switch and for power adjustment</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Pockels cell driver</td>
			<td>Conoptics</td>
			<td>M302-RM</td>
			<td>High voltage differential amplifier</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Shutter</td>
			<td>Vincent Associates</td>
			<td>LS6ZM2</td>
			<td>ALMgF2-coated BeCu blades</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Shutter controller</td>
			<td>Custom built</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Shutter driver</td>
			<td>Vincent Associates</td>
			<td>Uniblitz VCM D1</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Slicer / Vibratome</td>
			<td>Thermo Scientific</td>
			<td>Microm HM 650 V</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Uncaging software</td>
			<td>Rapp OptoElectronic</td>
			<td>UGA40 1.1.2.6 + NetCam</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">UV laser&#160;</td>
			<td>Rapp OptoElectronic</td>
			<td>DL-355/10</td>
			<td>cw DPSSL Laser, 355 nm; 10 mW; CAUTION</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">UV laser scanning system</td>
			<td>Rapp OptoElectronic</td>
			<td>UGA 40</td>
			<td>Galvanometric scanning system</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Name of Compound</td>
			<td>Catalog Number</td>
			<td></td>
			<td>Comments/Description</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">CNQX</td>
			<td>Sigma Aldrich</td>
			<td>C-127</td>
			<td>AMPA receptor antagonist; CAUTION</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">DL-AP5</td>
			<td>Alfa Aesar</td>
			<td>J64210</td>
			<td>NMDA receptor antagonist; CAUTION</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">SBFI K+ Salt</td>
			<td>Teflabs</td>
			<td>OO32</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">TTX</td>
			<td>Ascent Scientific</td>
			<td>Asc-055</td>
			<td>Inhibitor of voltage-dependent Na+ channels; CAUTION</td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">NMI-caged-L-Glutamate</td>
			<td>Torcis</td>
			<td>1490</td>
			<td>Caged glutamatereleased upon UV absobtion</td>
		</tr>
	</tbody>
</table>

multi-photon intracellular sodium imaging combined with uv-mediated focal uncaging of glutamate in ca1 pyramidal neurons

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high spatial and temporal resolution. Combined with electrophysiology, it is widely used to study activity-related calcium signals in small subcellular compartments such as dendrites and dendritic spines. In addition to calcium transients, synaptic activity also induces postsynaptic sodium signals, the properties of which are only marginally understood. Here, we describe a method for combined whole-cell patch-clamp and multi-photon sodium imaging in cellular micro domains of central neurons. Furthermore, we introduce a modified procedure for ultra-violet (UV)-light-induced uncaging of glutamate, which allows reliable and focal activation of glutamate receptors in the tissue. To this end, whole-cell recordings were performed on Cornu Ammonis subdivision 1 (CA1) pyramidal neurons in acute tissue slices of the mouse hippocampus. Neurons were filled with the sodium-sensitive fluorescent dye SBFI through the patch-pipette, and multi-photon excitation of SBFI enabled the visualization of dendrites and adjacent spines. To establish UV-induced focal uncaging, several parameters including light intensity, volume affected by the UV uncaging beam, positioning of the beam as well as concentration of the caged compound were tested and optimized. Our results show that local perfusion with caged glutamate (MNI-Glutamate) and its focal UV-uncaging result in inward currents and sodium transients in dendrites and spines. Time course and amplitude of both inward currents and sodium signals correlate with the duration of the uncaging pulse. Furthermore, our results show that intracellular sodium signals are blocked in the presence of blockers for ionotropic glutamate receptors, demonstrating that they are mediated by sodium influx though this pathway. In summary, our method provides a reliable tool for the investigation of intracellular sodium signals induced by focal receptor activation in intact brain tissue.

In the present study, we demonstrate a procedure for multi-photon microscopy of cellular sodium dynamics with the sodium-dependent fluorescent dye SBFI in CA1 pyramidal neurons of acute mouse hippocampal tissue slices. Moreover, we show how to combine this imaging technique with laser-scanning-based uncaging of neuro-active compounds (e.g. caged glutamate) and its precise targeting to cellular micro-domains.
Loading neurons with SBFI through the patch-pipette enabled the visualization of the entire cell including fine dendrites and adjacent spines by employing multi-photon excitation (Figure 6A, B, D and Figure 7A).
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/52038/52038fig6highres.jpg" width="500" /> 
Figure 6. Sodium signals and synaptic currents induced by flash photolysis of caged glutamate. (A) Maximal projection image of a CA1 pyramidal neuron loaded with SBFI via the patch pipette (PP). The box indicates the area shown enlarged in B. LP indicates the position and orientation of the pipette for local perfusion of caged glutamate. (B) High power maximum projection of a stack of optical sections of a dendrite with adjacent dendritic spines. The stacks of optical sections underwent z-alignment and deconvolution. The red cross indicates the target region of the uncaging beam. The orange dotted line delineates the region of interest from which the fluorescence emission was recorded. The box indicates the area shown enlarged in D. (C) Left: Sodium signal (upper row) and somatic inward current (lower row) induced by flash photolysis of caged glutamate (indicated by yellow flash). The red line represents a fit of the experimental data. The grey area represents the period in which the uncaging flash (300 msec) obstructed the imaging of SBFI fluorescence. Right: Without pre-perfusion with caged glutamate, the same UV-flash neither evoked a change in SBFI emission, nor an inward current. (D) Left: High power image of the dendrite and adjacent spines as delineated in B. Alongside, the same image with inverted grey values. The dotted lines indicate the regions of interest in which the fluorescence emission was recorded. The red cross indicates the localization of the uncaging beam. Right: Sodium transients induced by UV-flash photolysis of caged glutamate. Blue trace: sodium signals in the dendrite; red trace: averaged response from three spines adjacent to the uncaging spot; green trace: averaged response from three distant spines. <a href="https://www.jove.com/files/ftp_upload/52038/52038fig6highres.jpg" target="_blank">Please click here for a larger version of this figure.</a>
After local perfusion with caged glutamate, applying a UV-flash close to a dendrite resulted in a transient decrease in fluorescence emission of SBFI, reflecting an increase in the intracellular sodium concentration (Figure 6C, D and Figure 7B). At the same time, an inward current was recorded at the soma (Figure 6C and Figure 7B). Increasing the duration of the UV flash resulted in increasing amplitudes of both the elicited inward currents and the sodium signals (data not shown), indicating that the system was well within its dynamic range and that neither uncaging nor cellular responses were saturated. Application of a UV-flash of identical or longer duration to slices which had not been pre-perfused with caged glutamate, never elicited changes in SBFI fluorescence nor inward currents, indicating that these signals are due to the uncaging of glutamate (Figure 6C). Moreover, these results show that no interdependency of imaging- and uncaging components is observed under our experimental conditions. Sodium signals could also be detected in dendritic spines (Figure 6D). While we did not attempt to achieve stimulation of a single spine only with glutamate uncaging, peak amplitudes tended to be slightly higher in spines close to the uncaging spot, whereas spines further away showed virtually identical fluorescence changes as the parent dendrite (Figure 6D).
Finally, we studied the pathway for sodium influx into dendrites and spines in response to uncaging of glutamate. To this end, we employed CNQX and APV, which are selective blockers for the sodium-permeable, ionotropic glutamate receptors of the AMPA- and NMDA-subtype, respectively. Our results show that glutamate-induced intracellular sodium signals and the elicited somatic currents were omitted in the presence of these blockers (Figure 7B). Upon wash-out of the blockers, the signals are regained. This demonstrates that uncaging of glutamate activates ionotropic glutamate receptors on CA1 pyramidal neurons, which mediate the influx of sodium into dendrites and spines, resulting in intracellular sodium transients and inward currents.
<img alt="Figure 7" fo:content-width="6in" src="/files/ftp_upload/52038/52038fig7highres.jpg" width="600" /> 
Figure 7. Pharmacological profile of evoked sodium signals and inward currents. (A) SBFI-filled CA1 pyramidal neuron with patch pipette (PP) attached to the soma. (B) Left: Sodium transient and somatic current induced by photo-activation of caged glutamate in a dendrite and attached spines. Center: Perfusion with the ionotropic glutamate receptor blockers CNQX and APV inhibits both the sodium signal and the inward current induced by uncaging. Right: Upon wash-out of the blockers, the signals are restored. The red line represents a fit of the experimental data. The grey area represents the period in which the uncaging flash (300 msec) obstructed the imaging of SBFI fluorescence. <a href="https://www.jove.com/files/ftp_upload/52038/52038fig7highres.jpg" target="_blank">Please click here for a larger version of this figure.</a>

Watch this Scientific Journal Video about Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons at JoVE.com

Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

We describe the combination of focal UV-induced photo-activation of neuro-active compounds with whole-cell patch-clamp and multi-photon imaging of intracellular sodium transients in dendrites and spines of hippocampal neurons in acute tissue slices of the mouse brain.

Multi-photon fluorescence microscopy has enabled the analysis of morphological and physiological parameters of brain cells in the intact tissue with high ...

multi-photon-intracellular-sodium-imaging-combined-with-uv-mediated

Research

JoVE Journal

Neuroscience

14.0K 조회수.  Heinrich Heine University Düsseldorf.  우리는 전체 세포 패치 - 클램프 및 마우스 뇌의 급성 조직 슬라이스에서 수상 돌기 및 해마 신경 세포의 쪽의 세포 내 나트륨 과도 다중 광자 이미징 신경 활성 화합물의 초점 UV 유도 된 사진 활성화의 조합을 설명합니다. 

비디오: Multi-photon Intracellular Sodium Imaging Combined with UV-mediated Focal Uncaging of Glutamate in CA1 Pyramidal Neurons

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

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

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

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

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

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

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

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

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

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

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

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

Functional Interrogation of Adult Hypothalamic Neurogenesis with Focal Radiological Inhibition

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral delivery or transgenic animals have potential confounds that make interpretation of results from these studies difficult. New radiological tools are emerging, however, that allow one to noninvasively investigate the function of select groups of adult-born neurons through accurate and precise anatomical targeting in small animals. Focal ionizing radiation inhibits the birth and differentiation of new neurons, and allows targeting of specific neural progenitor regions. In order to illuminate the potential functional role that adult hypothalamic neurogenesis plays in the regulation of physiological processes, we developed a noninvasive focal irradiation technique to selectively inhibit the birth of adult-born neurons in the hypothalamic median eminence. We describe a method for Computer tomography-guided focal irradiation (CFIR) delivery to enable precise and accurate anatomical targeting in small animals. CFIR uses three-dimensional volumetric image guidance for localization and targeting of the radiation dose, minimizes radiation exposure to nontargeted brain regions, and allows for conformal dose distribution with sharp beam boundaries. This protocol allows one to ask questions regarding the function of adult-born neurons, but also opens areas to questions in areas of radiobiology, tumor biology, and immunology. These radiological tools will facilitate the translation of discoveries at the bench to the bedside.

The function of adult-born mammalian neurons remains an active area of investigation. Ionizing radiation inhibits the birth of new neurons. Using computer tomography-guided focal irradiation (CFIR), three-dimensional anatomical targeting of specific neural progenitor populations can now be used to assess the functional role of adult neurogenesis.

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral ...

Olfactory Neurons Obtained through Nasal Biopsy Combined with Laser-Capture Microdissection: A Potential Approach to Study Treatment Response in Mental Disorders

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium carbonate. Animal studies as well as human genetic studies indicate that lithium affects molecular targets that are involved in neuronal growth, survival and maturation, and notably molecules involved in Wnt signaling. Given the ethical challenge to obtaining brain biopsies for investigating dynamic molecular changes associated with lithium-response in the central nervous system (CNS), one may consider the use of neurons obtained from olfactory tissues to achieve this goal.The olfactory epithelium contains olfactory receptor neurons at different stages of development and glial-like supporting cells. This provides a unique opportunity to study dynamic changes in the CNS of patients with neuropsychiatric diseases, using olfactory tissue safely obtained from nasal biopsies. To overcome the drawback posed by substantial contamination of biopsied olfactory tissue with non-neuronal cells, a novel approach to obtain enriched neuronal cell populations was developed by combining nasal biopsies with laser-capture microdissection. In this study, a system for investigating treatment-associated dynamic molecular changes in neuronal tissue was developed and validated, using a small pilot sample of BD patients recruited for the study of the molecular mechanisms of lithium treatment response.

In this study, a novel platform to investigate intraneuronal molecular signatures of treatment response in bipolar disorder (BD) was developed and validated. Olfactory epithelium from BD patients was obtained through nasal biopsies. Then laser-capture microdissection was combined with Real Time RT PCR to investigate the molecular signature of lithium response in BD.

Bipolar disorder (BD) is a severe neuropsychiatric disorder with poorly understood pathophysiology and typically treated with the mood stabilizer, lithium ...

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

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

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

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

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

Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate Receptors in the Mouse Amygdala

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to study the molecular composition of membranes produced a significant decline in its use. Recently, there has been a major revival in freeze-fracture electron microscopy thanks to the development of effective ways to reveal integral membrane proteins by immunogold labeling. One of these methods is known as detergent-solubilized Freeze-fracture Replica Immunolabeling (FRIL).
The combination of the FRIL technique with optogenetics allows a correlated analysis of the structural and functional properties of central synapses. Using this approach it is possible to identify and characterize both pre- and postsynaptic neurons by their respective expression of a tagged channelrhodopsin and specific molecular markers. The distinctive appearance of the postsynaptic membrane specialization of glutamatergic synapses further allows, upon labeling of ionotropic glutamate receptors, to quantify and analyze the intrasynaptic distribution of these receptors. Here, we give a step-by-step description of the procedures required to prepare paired replicas and how to immunolabel them. We will also discuss the caveats and limitations of the FRIL technique, in particular those associated with potential sampling biases. The high reproducibility and versatility of the FRIL technique, when combined with optogenetics, offers a very powerful approach for the characterization of different aspects of synaptic transmission at identified neuronal microcircuits in the brain.
Here, we provide an example how this approach was used to gain insights into structure-function relationships of excitatory synapses at neurons of the intercalated cell masses of the mouse amygdala. In particular, we have investigated the expression of ionotropic glutamate receptors at identified inputs originated from the thalamic posterior intralaminar and medial geniculate nuclei. These synapses were shown to relay sensory information relevant for fear learning and to undergo plastic changes upon fear conditioning.

This article illustrates how the expression of neurotransmitter receptors can be quantified and the pattern analyzed at synapses with identified pre and postsynaptic elements using a combination of viral transduction of optogenetic tools and the freeze-fracture replica immunolabeling technique.

Freeze-fracture electron microscopy has been a major technique in ultrastructural research for over 40 years. However, the lack of effective means to ...

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 ...