This research was supported by grants from the LuMind Foundation, Research Down Syndrome, and the Alzheimer&#8217;s Association (AS). CP was partially supported by a faculty development grant from the College of Nursing and Health Professions at Arkansas State University.

NOTE: Experiments were conducted in accordance with the ethical standards approved by the Committee on Animal Research at the Veterans Affairs Palo Alto Health Care System.
1. Golgi-Cox Staining
<ol>
	<li>Brain extraction and staining
	<ol>
		<li>On the 1st day, deeply anesthetize mice with 100 mg/kg ketamine and 10 mg/kg xylazine before euthanizing via exsanguination.</li>
		<li>Carefully remove the calvarium and dissect out the brain.
		<ol>
			<li>First remove the skin on the top of the skull, placing a curved scissor on the top of the cerebellum and gently cut through the calvarium parallel to the central sulcus with the tip of the scissor pointed outward, moving in the direction toward the olfactory bulb. Repeat this process on both sides.</li>
			<li>Use a surgical pincet to lift the calvarium while turning it toward the olfactory bulbs and break it off. Then, flip the skull upside down and use a pick to cut through the optic nerves and loosen the brain. Finally, use the pick to separate the brain from the spinal cord at the level of the foramen magnum.</li>
		</ol>
		</li>
		<li>Immediately immerse the brain in 15 ml of commercially-available, undiluted Golgi-Cox solution. Store the brain in Golgi solution in a capped, 20 ml glass bottle at room temperature in the dark.</li>
		<li>On the 2nd day, slowly pour out the solution and replace it with 15 ml fresh Golgi-Cox solution. Recap the bottle containing the brains and leave undisturbed for another 9 days in the dark.</li>
	</ol>
	</li>
	<li>Sectioning and embedment
	<ol>
		<li>On the 11th day, place the brains in 30% sucrose solution, prepared in ddH2O for dehydration. Change the solution with freshly prepared 30% sucrose after 12 hours. Incubate the brains in a 30% sucrose solution for 3 days at 4 &#176;C.</li>
		<li>On the 14th day, fill the reservoir of the vibratome with a 30% sucrose solution until the blade is covered. Set the speed to 1 mm/s with amplitude of 0.95 mm.</li>
		<li>Blot the brains with a Kimwipe to remove excess moisture and remove the cerebellum using a razor blade to cut coronally.</li>
		<li>Firmly mount the whole brain onto the vibratome platform using superglue and allow it to set for 2-4 min. Insert the platform with the adhered brain into the reservoir and adjust the blade to the top of the brain.</li>
		<li>Using the vibratome, slice the brain into 150 &#181;m thick sections at room temperature, remembering to change the blade after every 3rd brain.</li>
		<li>Pick up the sections from the reservoir using a small tipped brush and immerse them into &#160;a petri dish containing 0.3% gelatin made in ddH2O. While the sections remain in the petri dish, add 0.5 ml of 0.3% gelatin on each slide and use a brush to spread and coat the superfrost + slides.</li>
		<li>Gently pick up the immersed sections from the petri dish and place them onto the gelatinized slides. Orient the brain sections by touching them only on their sides and not on the tops.</li>
		<li>After about ten min, coat the sections with 30% sucrose before the tissue completely dries. Then air dry the prepared slides in a dark place in a slide rack for 3 days.</li>
		<li>Dilute commercially-available 10x Developing solution to 1x using ddH2O. Immerse the slides in developing solution for 7-10 min. Rinse the slides 3 times in ddH2O.</li>
	</ol>
	</li>
	<li>Dehydration
	<ol>
		<li>Prepare 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, and 95% ethanol solutions using distilled ddH2O.</li>
		<li>Soak the slides in increasingly higher ethanol solutions for 10 min each.</li>
		<li>Soak the slides in 100% ethanol solution 3 times for 10 min each time.</li>
		<li>Soak the slides in 100% xylene 3 consecutive times for 5 min each, keeping the sections in the final solution until they are cover-slipped.</li>
		<li>Place a generous amount of DPX (Di-n-butyl phthalate in xylene) mounting medium (about 1-1.5 ml) on each slide using a plastic transfer pipet. Carefully place coverslips to avoid bubbles.</li>
		<li>Air dry cover slipped-brain sections horizontally for 3 days.</li>
	</ol>
	</li>
</ol>
2. Doublecortin Staining
<ol>
	<li>Deeply anesthetize (see step 1.1.1) the animals.</li>
	<li>Perform cardiac perfusion with freshly prepared 4% paraformaldehyde made in phosphate buffer12.</li>
	<li>Extract the brains (see 1.1.2) and store in 45 ml of 4% paraformaldehyde at 4 &#176;C overnight on a rocking shaker.</li>
	<li>Transfer the brains to a 30% sucrose solution and wait for complete dehydration for 48 hr at 4 &#176;C. Remove the brains from sucrose and place them directly on copper blocks placed on dry ice.</li>
	<li>Fill the block with optimum cutting temperature (OCT) compound and mark the orientation of the brain using the olfactory bulb as a landmark.</li>
	<li>Using a cryostat, cut 70 micron-thick sections at -20 &#186;C and place them in cryoprotectant solution (25% ethylene glycol, 25% glycerol, in 0.05 M sodium phosphate buffer, pH 7.4) and keep at -20 &#176;C until use.</li>
	<li>Make a solution of 50% cryoprotectant and 50% methanol. To this solution add 0.5% hydrogen peroxide (vol/vol). Incubate floating sections for 30 min at room temperature.</li>
	<li>Warm sections in 37 &#186;C in TBS (Tris-buffered saline) for 30-40 min by placing them in a histological oven.</li>
	<li>Incubate the sections with 0.3% triton and 3% normal horse serum for 45 min at room temperature prior to immunostaining. Incubate the sections at 4 &#186;C overnight in 3 ml solution of the DCX antibody diluted 1:500 in TBS with 0.3% triton and 1% normal horse serum.</li>
	<li>The following day wash the sections 3 consecutive times in TBS (10 min each).</li>
	<li>Incubate sections with a biotinylated horse anti-goat (1:200 in TBS) for two hr at room temperature. Wash the sections 3 consecutive times in TBS (10 min each).</li>
	<li>Incubate them with ABC Lite (1:1,000) for 1.5 hr at room temperature.</li>
	<li>Add 5 &#181;l of 30% H2O2 to one tablet of 10 mg of DAB (3,3&#8221;-Diaminobenzidine) solution dissolved in 15 ml of 1 M Tris-HCl (pH 7.6). Incubate the sections immediately in this solution for 5 min.</li>
	<li>Terminate the reaction by washing them with ice-cold TBS three times followed by one wash in TBS at room temperature.</li>
	<li>Dehydrate sections using a series of ethanol solutions (50%, 60%, 70%, 80%, 90%, 95%, 100%, 5 min each), clear in xylene, and then cover slip using DPX as in step 1.3.</li>
</ol>
3. Visualization
<ol>
	<li>Following complete drying of the sections, remove&#160;excess DPX on top of the slides with a sharp blade, and place them on the scanning stage of the microscope. The system is composed of a microscope equipped with scanning stage, joystick, and a color 12-bit camera.</li>
	<li>Start the program. Open a new data file.</li>
	<li>Place a reference point on the screen by clicking with the mouse pointer at any place. This will activate all the icons from the tool panel of program window.</li>
	<li>Click on the &#8220;Joystick Free&#8221; icon on the toolbar and then use the joystick to locate the dentate gyrus of the hippocampus in the first section.</li>
	<li>In the &#8220;Tools&#8221; tab of the program window, select &#8220;Serial Section Manager&#8221;. Click on the &#8220;New Section&#8221; icon on the lower left corner of the window. This will open the &#8220;Serial Section Setup&#8221; window.</li>
	<li>Select the Serial Section Setup window and then enter the total number of sections that contain hippocampal regions. Select the evaluation interval. Enter the section cut thickness (70 &#181;m for DCX and 150 &#181;m for Golgi-stained sections).</li>
	<li>Start tracing the dentate gyrus region by clicking on the &#8220;Freehand Contour Drawing&#8221; icon in the tool bar. Outline the dentate granular cell layer.</li>
	<li>Select &#8220;100X&#8221; from the &#34;Magnification&#34; menu. Add a drop of immersion oil on the section and switch the objective to 100X. Locate the dentate granule cell bodies and dendrites under this objective.</li>
	<li>Focus on a selected neuron and click on the &#8220;Neuron Tracing&#8221; icon on the toolbar. Trace the circumference of the dentate granule cell body. When finished tracing, right click to select &#8220;Finish Cell Body&#8221;.</li>
	<li>Start tracing the dendrites manually in x, y, and z directions and follow each branch using the joystick and z-motor knob. At a bifurcation or trifurcation node, select the respective option from the dropdown menu upon right-click. Trace each of the branches that arise from these nodes. At the end of each branch right click and select &#8220;Ending&#8221; from the dropdown menu.</li>
	<li>Using the arrow key icons in the software window randomly select another dentate granule cell and follow the same procedure as described in step 3.9-3.10. Save the entire tracing.</li>
	<li>Start the neuronal tracing software.</li>
	<li>Open the first NRX data file from the first mouse of the experimental group. Append the NRX file of the second mouse of the experimental group. Continue until all the NRX files from this group are appended.</li>
	<li>Under the Analysis tab, select &#8220;Fan In-diagram&#8221;. This opens the Fan In-analysis window. Check mark &#8220;dendrites&#8221; and click Display, which depicts the lengths and branching patterns of dendrites for this group of mice (Figure 1).</li>
</ol>
4. EDFI
NOTE: This method enables the user to rapidly record a large number of images through the z-axis of each field and generate a 2D image that contains all the focused pixels throughout the z-axis. The result will be a composite image that contains all focused pixels from collected images.
<ol>
	<li>Connect the microscope to a digital camera. Place the sections first on the scanning stage connected to the microscope and then switch to a 10X objective. Move the stage to the area of interest.</li>
	<li>Start a video capturing program.</li>
	<li>Press the record button. As soon as the record button is pressed, use the macro-focus knob to move from the top to the bottom of the section for a total of 4 sec. Save the resulting video file.</li>
	<li>Use ImageJ freeware to convert the AVI video files into an uncompressed format by opening the file and saving them in uncompressed file format.</li>
	<li>Start an image analysis program and open the AVI file. Go to &#8220;Process&#8221; menu and click on &#8220;Extended Depth of Field&#8221;. Select the corresponding video file. In &#8220;Output&#8221; options, select &#8220;Generate Composite Best-Focus Image&#8221;.</li>
	<li>In &#8220;Focus Analysis&#8221; options, select &#8220;Normalized Illumination&#8221; and &#8220;Max Local Contrast&#8221; options. Then, select &#8220;Create&#8221;. The resulting image represents all the focused pixels throughout the z-axis (Figure 2).</li>
	<li>Save the resulting image.</li>
</ol>
5. Generating Extremely-high Resolution Images
NOTE: This method allows the user to automatically generate low magnification and extremely high-resolution images from high-magnification-high resolution images in an automatized fashion.
<ol>
	<li>Place the sections first on the scanning stage connected to the microscope. The microscope is connected to a digital camera. Move the stage to the area of interest.</li>
	<li>Start an image acquisition program.</li>
	<li>Use a 10X objective and acquire and save high-resolution (4076 x 3116) images from the region of interest making sure to have at least 10% overlap.</li>
	<li>Run Image Composite Editor to stitch images.</li>
	<li>Go to File menu, and click on &#8220;New Panorama&#8221;. Select the folder in which the images are stored. Select the images that belong to the same region and press &#8220;ok&#8221;.</li>
	<li>Press &#8220;Export to Disk&#8221; button and save the image. This image represents an extremely high-resolution picture of the area of interest that could be used for analysis and/or demonstration (Figure 3).</li>
</ol>

<ol>
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Publication fees for this article are sponsored by MBF Bioscience.

Here, two methods were described to quantify the extent of dendritic arborization in mature and newly-born neurons using conventional staining methods in conjunction with RTI and EDFI. The acquisition of high resolution images of neurons provides an extremely useful method for testing the deleterious effects of neurodegenerative disorders and, in turn, provides a means to assess therapeutic strategies that target hippocampal neurons.
While the RTI method was utilized to capture in-depth data p...

Dynamic alterations in the number and structure of synapses are hallmarks of development, aging, and numerous neurodegenerative disorders1-3. The ability of neurons to receive and integrate synaptic information depends upon dendritic morphology and dynamic alterations in synaptic connections. Indeed, a positive correlation exists between dendritic spine and synapse number, which both impact cognitive function4. Thus, it is not surprising that decrements in dendritic spine number have been associated with cognitive dysfunction in a number of neurological disorders5-7, prompting great interest in dendritic spine quantification. Nevertheless, the quantification of spine density remains a time-consuming and tedious task that fails to generate useful information regarding the topography and distribution of synapses across the dendritic tree. Fortunately, staining methods (e.g., Golgi-Cox and doublecortin (DCX)) in conjunction with sophisticated imaging techniques can be utilized to overcome current barriers and produce high-resolution images of dendritic arborization in a reliable and expeditious manner. While Golgi-Cox staining method can be deployed to assess the state of dendritic arborization in all neurons8, DCX can be deployed to label newly-born neurons particularly in the dentate gyrus and subventricular zone9, an important consideration given that neurogenesis occurs in both these regions throughout the lifespan10,11.
Following staining, two imaging methods were deployed to assess dendritic characteristics: i) real-time imaging (RTI) and ii) extended depth of field imaging (EDFI). The RTI technique provides a mean to trace and quantify the length and order of arborization along the individual dendritic segments and branches. Thus it enables one to estimate the total area and volume occupied by each dendritic tree. More specifically, in the RTI method the user continuously identifies the segments and refocuses iteratively as the neuron tracing software collects the x, y, and z coordinates of the dendritic structure and reconstructs the trajectory of the dendritic structure in 3D. Comparatively, the EDFI method provides a rather simple and expedited means for assessing dendritic density in rather thick tissue specimens by generating a composite image, providing information on the entire z-axis. To do so, the user records high definition video files throughout the thickness of the section and then uses software to search the video frames to identify points wherein a pixel is completely in focus. Subsequently, the focused pixels are merged and integrated into a high-resolution, composite 2D image. This composite image contains all pixels that were in-focus regardless of their position in the z axis. Qualitative and quantitative analysis of these 2D images can be used subsequently to determine the density of dendritic branching in each field.
Lastly, we present a panoramic method for generating extremely high-resolution images for the analysis and assessment of dendrites in an entire region of interest. This technique can be deployed to overcome the lack of access to very high-resolution and expensive digital cameras. Using this method, one captures serial images at different locations along the x- and y-axes and then automatically stitches them together using a freeware (e.g., Image Composite Editor). Notably, this method can be used for qualitative and quantitative assessment of dendritic arborization in a rather wide area.

<table border="0" cellpadding="0" cellspacing="0" width="960">
	<tbody>
		<tr height="25">
			<td height="25" style="height:25px;width:272px;">Name</td>
			<td style="width:263px;">Company</td>
			<td style="width:267px;">Catalog Number</td>
			<td style="width:159px;">Comments</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">Modified Golgi-cox staining solution&#160;</td>
			<td style="width:263px;">Weill Cornell Medical College</td>
			<td style="width:267px;">NA</td>
			<td style="width:159px;">store at 4&#176;C till use</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">1x Developing Solution (Stock 10x)</td>
			<td style="width:263px;">Weill Cornell Medical College</td>
			<td style="width:267px;">NA</td>
			<td style="width:159px;">store at 4&#176;C till use</td>
		</tr>
		<tr height="26">
			<td height="26" style="height:26px;width:272px;">30% Sucrose,</td>
			<td style="width:263px;">Sigma</td>
			<td style="width:267px;">CAS # 57-50-1</td>
			<td style="width:159px;">make fresh &#160;in ddH2O</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">0.3% Gelatin</td>
			<td style="width:263px;">Sigma</td>
			<td style="width:267px;">CAS # 9000-70-8</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:272px;">Graded Ethanol Solutions (20%, 30%, 40%, 50%, 80%. 90%, 95%. 100%)</td>
			<td style="width:263px;">Sigma</td>
			<td style="width:267px;">CAS 603-003-00-5</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">Xylene</td>
			<td style="width:263px;">Sigma</td>
			<td style="width:267px;">CAS # 1330-20-7</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">DPX Medium</td>
			<td style="width:263px;">EMS&#160;</td>
			<td style="width:267px;">#13510</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">Superfrost (+) white</td>
			<td style="width:263px;">Electron Microscopy Sciences</td>
			<td style="width:267px;">71869-10</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">Coverslip 22x50mm (VWR #48393-059)</td>
			<td style="width:263px;">VWR&#160;</td>
			<td style="width:267px;">#4811-703</td>
			<td style="width:159px;">NA</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">DCX Antibody</td>
			<td style="width:263px;">Santa Cruz Biotechnology</td>
			<td style="width:267px;">sc-8066</td>
			<td style="width:159px;">4 C</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">DAB</td>
			<td style="width:263px;">Sigma</td>
			<td style="width:267px;">CAS Number 91-95-2 &#160;</td>
			<td style="width:159px;">-20</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:272px;">OCT</td>
			<td style="width:263px;">Tissue-tek</td>
			<td style="width:267px;">4583</td>
			<td style="width:159px;">NA</td>
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assessment of dendritic arborization in the dentate gyrus of the hippocampal region in mice

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity and extent of dendritic arborization correlates well with the synaptic plasticity in these cells. A reliable method for assessment of dendritic arborization is needed to characterize the deleterious effects of neurological disorders on these structures and to determine the effects of therapeutic interventions. However, quantification of these structures has proven to be a formidable task given their complex and dynamic nature. Fortunately, sophisticated imaging techniques can be paired with conventional staining methods to assess the state of dendritic arborization, providing a more reliable and expeditious means of assessment. Below is an example of how these imaging techniques were paired with staining methods to characterize the dendritic arborization in wild type mice. These complementary imaging methods can be used to qualitatively and quantitatively assess dendritic arborization that span a rather wide area within the hippocampal region.

The extent of arborization arising from extant and newly born dentate granule cells was analyzed in wild type mice using either Golgi-Cox or DCX staining (Figure 1). Dendritic segments of DCX-positive cells were found to be 13-36 microns long. The normal distribution of dendritic length was tested using Kolmogorov-Smirnov test (D = 0.1217, p &#60;0.01, Liliefors p &#60;0.001; Figures 4 and 5).
In the analysis of length of segments per order of...

Watch this Scientific Journal Video about Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice at JoVE.com

Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

We describe two methods for visualization and quantification of dendritic arborization in the hippocampus of mouse models: real-time and extended depth of field imaging. While the former method allows sophisticated topographical tracing and quantification of the extent of branching, the latter allows speedy visualization of the dendritic tree.

Dendritic arborization has been shown to be a reliable marker for examination of structural and functional integrity of neurons. Indeed, the complexity ...

assessment-dendritic-arborization-dentate-gyrus-hippocampal-region

Research

JoVE Journal

Neuroscience

10.2K Views.  VA Palo Alto Health Care System. We describe two methods for visualization and quantification of dendritic arborization in the hippocampus of mouse models: real-time and extended depth of field imaging. While the former method allows sophisticated topographical tracing and quantification of the extent of branching, the latter allows speedy visualization of the dendritic tree.

Video: Assessment of Dendritic Arborization in the Dentate Gyrus of the Hippocampal Region in Mice

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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 ...

The Use of Trace Eyeblink Classical Conditioning to Assess Hippocampal Dysfunction in a Rat Model of Fetal Alcohol Spectrum Disorders

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain undergoes rapid organizational change and is similar to accelerated brain changes that occur during the third trimester in humans. This model of fetal alcohol spectrum disorders (FASDs) produces severe brain damage, mimicking the amount and pattern of binge-drinking that occurs in some pregnant alcoholic mothers. We describe the use of trace eyeblink classical conditioning (ECC), a higher-order variant of associative learning, to assess long-term hippocampal dysfunction that is typically seen in alcohol-exposed adult offspring. At 90 days of age, rodents were surgically prepared with recording and stimulating electrodes, which measured electromyographic (EMG) blink activity from the left eyelid muscle and delivered mild shock posterior to the left eye, respectively. After a 5 day recovery period, they underwent 6 sessions of trace ECC to determine associative learning differences between alcohol-exposed and control rats. Trace ECC is one of many possible ECC procedures that can be easily modified using the same equipment and software, so that different neural systems can be assessed. ECC procedures in general, can be used as diagnostic tools for detecting neural pathology in different brain systems and different conditions that insult the brain.

Trace eyeblink classical conditioning (ECC) was used to assess hippocampal-dependent associative learning in adult rats that were administered a high concentration (11.9% v/v) of alcohol during early neonatal brain development. In general, ECC procedures are sound diagnostic tools for detecting brain dysfunction across many psychological and biomedical settings.

Neonatal rats were administered a relatively high concentration of ethyl alcohol (11.9% v/v) during postnatal days 4-9, a time when the fetal brain ...

Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation. There are several approaches to Golgi staining, all of which have been useful for determining the morphological characteristics of dendritic arbors and produce a clear background. The present Golgi-Cox method, (a slight variation of the protocol that is provided with a commercial Golgi staining kit), was designed to assess how a relatively low dose of the chemotherapeutic drug 5-flurouracil (5-Fu) would affect dendritic morphology, the number of spines, and the complexity of arborization within the hippocampus. The 5-Fu significantly modulated the dendritic complexity and decreased the spine density throughout the hippocampus in a region-specific manner. The data presented show that the Golgi staining method effectively stained the mature neurons in the CA1, the CA3, and the dentate gyrus (DG) of the hippocampus. This protocol reports the details for each step so that other researchers can reliably stain tissue throughout the brain with high quality results and minimal troubleshooting.

Here we present a Golgi-Cox protocol in extensive detail. This reliable tissue stain method allows for a high-quality assessment of the cytoarchitecture in the hippocampus, and throughout the entire brain, with minimal troubleshooting.

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. Drosophila larval neuronal dendritic arborization (da) is an ideal model for studying morphogenesis of neural dendrites and gene function in the development of nervous system. There are four classes of da neurons. Class IV is the most complex with a branching pattern that covers almost the entire area of the larval body wall. We have previously characterized the effect of silencing the Drosophila ortholog of SOX5 on class IV neuronal dendritic arborization complexity (NDAC) using four parameters: the length of dendrites, the surface area of dendrite coverage, the total number of branches, and the branching structure. This protocol presents the workflow of NDAC quantitative analysis, consisting of larval dissection, confocal microscopy, and image analysis procedures using ImageJ software. Further insight into da neuronal development and its underlying mechanisms will improve the understanding of neuronal function and provide clues about the fundamental causes of neurological and neurodevelopmental disorders.

Quantitative Analysis of Neuronal Dendritic Arborization Complexity in Drosophila

This protocol focuses on quantitative analysis of neuronal dendritic arborization complexity (NDAC) in Drosophila, which can be used for studies of dendritic morphogenesis.

Dendrites are the branched projections of a neuron, and dendritic morphology reflects synaptic organization during the development of the nervous system. ...

An Invasive Method for the Activation of the Mouse Dentate Gyrus by High-frequency Stimulation

Electrical high-frequency stimulation (HFS), using implanted electrodes targeting various brain regions, has been proven as an effective treatment for various neurological and psychiatric disorders. HFS in the deep region of the brain, also named deep-brain stimulation (DBS), is becoming increasingly important in clinical trials. Recent progress in the field of high-frequency DBS (HF-DBS) surgery has begun to spread the possibility of utilizing this invasive technique to other situations, such as treatment for major depression disorder (MDD), obsessive-compulsive disorder (OCD), and so on. 
Despite these expanding indications, the underlying mechanisms of the beneficial effects of HF-DBS remain enigmatic. To address this question, one approach is to use implanted electrodes that sparsely activate distributed subpopulations of neurons by HFS. It has been reported that HFS in the anterior nucleus of the thalamus could be used for the treatment of refractory epilepsy in the clinic. The underlying mechanisms might be related to the increased neurogenesis and altered neuronal activity. Therefore, we are interested in exploring the physiological alterations by the detection of neuronal activity as well as neurogenesis in the mouse dentate gyrus (DG) before and after HFS treatment. 
In this manuscript, we describe methodologies for HFS to target the activation of the DG in mice, directly or indirectly and in an acute or chronic manner. In addition, we describe a detailed protocol for the preparation of brain slices for c-fos and Notch1 immunofluorescent staining to monitor the neuronal activity and signaling activation and for bromodeoxyuridine (BrdU) labeling to determine the neurogenesis after the HF-DBS induction. The activation of the neuronal activity and neurogenesis after the HF-DBS treatment provides direct neurobiological evidence and potential therapeutic benefits. Particularly, this methodology can be modified and applied to target other interested brain regions such as the basal ganglia and subthalamic regions for specific brain disorders in the clinic.

This protocol shows how to set up a reliable HFS method in mice. Neurons throughout the hippocampal dentate gyrus are electrically stimulated by HFS directly and indirectly in vivo. Neuronal activity and molecular signaling are examined by c-fos and Notch1 immunofluorescent staining, respectively; neurogenesis is quantified by bromodeoxyuridine labeling assay.

Electrical high-frequency stimulation (HFS), using implanted electrodes targeting various brain regions, has been proven as an effective treatment for ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.