This work was supported by a pilot grant under NIH P20 GM109005 (A.R.A.) and by the Center for Translational Neuroscience IDeA program award P30 GM110702.

Experiments were conducted in accordance with the ethical standards approved by the Institutional Animal Care and Use Committee at UAMS.
1. Animals and 5-Fu Injection Paradigm
<ol>
	<li>Purchase 6-month-old male C57Bl6/J wild-type mice and house them under a constant 12 h light/dark cycle until they reach 1 year of age.</li>
	<li>Dilute the 5-Fu in 0.9% sterile saline. Use 60 mg/kg as the required dose per mouse.</li>
	<li>Give the intraperitoneal injections of the 5-Fu (once per week for three weeks).
	<ol>
		<li>Give the intraperitoneal injections around the same time on each day. For example, between 0900-1200 h.</li>
	</ol>
	</li>
	<li>Euthanize the animals and extract the brains 30 days after the final injection.</li>
</ol>
2. Euthanasia Procedure and Brain Extraction
<ol>
	<li>Restrain the rodent and grab the base of the tail. With the other hand, place the thumb or first finger at the base of the rodent&#39;s neck. Quickly put pressure on the rodent&#39;s neck and push forward while the other hand holding the tail pulls backwards.</li>
	<li>Hold the rodent with one hand and the large surgical scissors with the other. Place the rodent&#39;s neck between the two blades and quickly decapitate the head.</li>
	<li>Take the severed mouse head, and use the fine scissors to trim the fur above the skull, up to the eyes. Next, place the tips of the scissors into each eye socket and close the scissors with slight force.</li>
	<li>Use spring scissors to cut the skull to the left and right of the cerebellum.</li>
	<li>Use forceps to lift the portion of the skull surrounding the cerebellum directly up and off.</li>
	<li>Use the spring scissors to cut along the midsagittal line of the skull.</li>
	<li>Use the forceps to lift the remaining portion of the skull off the brain.</li>
	<li>Use a spatula and a probe to scoop out the brain into the desired container. Using a stainless steel blade, cut each brain along the midsagittal plane. Perform the following Golgi-Cox method on the right hemispheres.</li>
</ol>
3. Golgi Staining and Tissue Preparation
<ol>
	<li>Immerse the freshly harvested half-brains into 5 mL mercuric chloride-based solution (solution A from the kit) in a 10 mL conical tube. The mercuric chloride solution is light-sensitive; cover the tube with foil. Store the sample in the dark at room temperature. 
	NOTE: Solution A is provided in the kit that is listed in the table of materials. Caution! Solution A (also called mercuric chloride solution) contains toxic reagents such as potassium dichromate, mercuric chloride, and potassium chromate. Wear protective gloves and work in a fume hood.</li>
	<li>After 1 day, slowly decant the solution into a temporary container (such as a large weigh boat), and dispose of it in a biohazard waste container. Immerse the brains (still in the original 10 mL conical tubes) with 5 mL of the mercuric chloride solution, and return the sample to the dark at room temperature for 13 more days.</li>
	<li>Add 30 g post-impregnation buffer to 90 mL distilled or deionized water (dH2O). Fill each well of a 6-well plate with 6.5 mL of the post-impregnation buffer, one well per brain 
	NOTE: Post-impregnation buffer is provided in the kit that is listed in the table of materials.
	<ol>
		<li>After 14 days (in total) in the mercuric chloride solution, rinse the tissue with dH2O, then transfer them into 6-well plates with post-impregnation buffer. Cover the plates with foil and store them at room temperature in the dark.</li>
	</ol>
	</li>
	<li>Pipette out the post-impregnation buffer after 1 day of immersion and renew with fresh post-impregnation buffer; store for 1 day in the dark at room temperature. If required, store this at 4 &#176;C for up to 1 month22.</li>
</ol>
4. Sectioning
<ol>
	<li>Label the 12-well plates on their lids with numbers in ascending order from left-to-right and up-to-down.</li>
	<li>Prepare two or three 12-well plates per brain if sectioning the entire brain.
	<ol>
		<li>Label the top of each 12-well plate lid with the identification number of the brain sectioned.</li>
		<li>Pipette 2 mL of 1x PBS into each well of each plate.</li>
	</ol>
	</li>
	<li>Set up the stage and turn on the vibratome; ensure that there is enough 1x PBS in the container to cover the tissue.
	<ol>
		<li>Cut two pieces of plastic paraffin film and fold them over twice. Place them over the holes for the specimen holder knobs to prevent 1x PBS from leaking onto the counter.</li>
		<li>Put tape down on the tissue block of the specimen holder and then put cyanoacrylate adhesive glue (see table of materials) on it.</li>
	</ol>
	</li>
	<li>Remove the brain from the 6-well plate and place it on a plastic or glass dish. Use the stainless steel double-edged blade to cut off about half of the cerebellum.
	<ol>
		<li>Cut the cerebellum caudally. Make sure that the portion of the cerebellum remaining is even.</li>
	</ol>
	</li>
	<li>Immediately place the flat surface of the remaining cerebellum on the glue. 
	NOTE: The dorsal part of the tissue (the cerebral cortex) should be facing left or right relative to the blade. The rostral end that previously contained the olfactory bulb attached should face upwards.
	<ol>
		<li>Wait at least a few minutes to make sure that the glue fully dries.</li>
	</ol>
	</li>
	<li>While the glue that is holding the brain in place is drying, pour 1x PBS into the specimen bath. After the glue is dry, place the specimen holder with the tissue into the specimen bath.
	<ol>
		<li>If the tissue sample is not fully covered, pour more 1x PBS into the specimen bath.</li>
	</ol>
	</li>
	<li>Attach the blade to the blade holder of the vibratome and slowly lower the blade into the specimen bath until it is fully submerged in 1x PBS. Continue lowering the blade until it is slightly below the top of the tissue.</li>
	<li>Set the vibratome speed to 7, the amplitude to 6, and the cutting angle to 12&#176; (see table of materials for vibratome model). Start the vibration and cut 200 &#181;m sections23.
	<ol>
		<li>Use a large paintbrush to move the tissue sections from the specimen bath into each designated well of the 12-well plates.</li>
	</ol>
	</li>
	<li>Continue to cut the tissue until the desired number of tissue sections is reached.</li>
</ol>
5. Post-staining
<ol>
	<li>For each of the following staining steps and rinses, use 2 mL of the indicated solution per well. Use a motorized pipet filler (see table of materials) to transfer the solutions into the wells. Use a transfer pipette (see table of materials) to transfer solutions out of the wells and into the proper waste containers.</li>
	<li>Rinse the sections in one 30 min wash of 0.01 M PBS-T (add 3.0 mL of detergent (see table of materials) to 1,000 mL 0.01 M PBS).</li>
	<li>Dilute ammonium hydroxide solution (Caution) 3:5 by volume with dH2O immediately before use. Stain the sections in the diluted ammonium hydroxide solution for 19-21 min.
	<ol>
		<li>Protect the light-sensitive ammonium hydroxide solution with foil. 
		NOTE: Ammonium hydroxide within solution has a concentration of ~10% and it is classified as &#34;hazardous.&#34; Ammonium hydroxide solution may produce burns if it contacts the skin. It may cause respiratory tract irritation if inhaled. Wear protective gloves and work in a fume hood.</li>
	</ol>
	</li>
	<li>Stain the sections in the post-staining buffer (add 90 g of reagent D in 500 mL dH2O) for 19-21 min. The post-staining buffer is light-sensitive; protect it with foil22.</li>
	<li>Rinse the sections in three, 5-10 min washes of 0.01 M PBS-T.</li>
</ol>
6. Mounting, Cleaning, and Covering
<ol>
	<li>Mount the sections on 1% gelatin-coated slides with the large paintbrush. Allow them to dry for 20-30 min at room temperature, and then place them in a Coplin jar overnight.</li>
	<li>Remove the slides with gloved hands from the Coplin jar and place them in a plastic rack (see table of materials). Place the plastic rack in the staining dish (see table of materials) containing 100% ethanol. Dehydrate them with three 5 min washes.</li>
	<li>Wash the slides twice for 5 min each in 99% xylene. Place the slides in a glass rack and lower the rack into a glass staining dish containing xylene with a metal handle (see table of materials). 
	NOTE: Caution, Xylene is harmful if inhaled and causes irritation if it touches the skin. It is highly flammable in liquid and vapor states. Wear protective gloves and work in a fume hood.</li>
	<li>Remove the slides from the xylene one at a time and quickly cover the tissue with ~0.25 mL of mounting media (see Material Table). Next, take the slide covers and lay them over the media.
	<ol>
		<li>Push any trapped air bubbles underneath the slide covers to the edge with a blunt object, such as the end of a paintbrush.</li>
	</ol>
	</li>
	<li>Place the slides in an area away from sunlight and allow them to dry for 2-3 days.</li>
	<li>Proceed to acquiring images using a microscope and analyzing with the appropriate software.</li>
</ol>
7. Acquire Image Stack
<ol>
	<li>Open the imaging program (see Materials Table). Place the slide onto the stage of the microscope. On the menu at the top of the program, click &#34;Acquisition&#34; and then click &#34;Live Image.&#34;</li>
	<li>Set the microscope to 10X. Identify the neuron of preference and then bring the cursor, which appears like a red X, onto the center of the soma. Left click to set the reference point.</li>
	<li>Set the microscope to 40X. Click &#34;Move&#34; from the menu at the top of the program and then click &#34;To Reference Point.&#34;</li>
	<li>Begin creating the image stack by manually scrolling with the fine objective above the tissue until it is slightly out of focus.
	<ol>
		<li>Click &#34;Set Top&#34; in the bottom right hand corner of the program under the heading &#34;Image Acquisition.&#34;</li>
		<li>Scroll slightly under the tissue until it is slightly out of focus and then click &#34;Set Bottom&#34; under &#34;Image Acquisition.&#34; Next, click &#34;Acquire Image Stack.&#34;</li>
	</ol>
	</li>
	<li>After the image stack is acquired, type the desired name for image file in the window that appears. Save as &#34;MBF Ascii file.&#34;
	<ol>
		<li>When prompted, select the file type as &#34;MBF JPEG2000 stack file&#34; and then click &#34;Save.&#34;</li>
	</ol>
	</li>
</ol>
8. Neuronal Tracing
<ol>
	<li>In the toolbar underneath the menu, click the box titled &#34;Contour Name.&#34; Select &#34;Soma&#34; from the drop down menu.</li>
	<li>Manually trace the soma. Then right click and select &#34;Finish Cell Body&#34; when tracing is complete.</li>
	<li>Select &#34;AutoNeuron&#34; to bring up another tab titled &#34;AutoNeuron Workflow.&#34;</li>
	<li>Under &#34;Configuration Type,&#34; select &#34;New.&#34; Set the parameters and then click &#34;Next Step&#34; to bring up &#34;Soma Detection&#34; subheading.
	<ol>
		<li>To set the parameters, select &#34;Brightfield&#34;. Next, under &#34;Max Process Diameter&#34;, click &#34;Start Measuring&#34;. Using the cursor, go to the base of the dendrite and manually set the diameter.</li>
	</ol>
	</li>
	<li>Click &#34;Yes&#34; in the window that requests the user to trace the entire image. Manually trace the soma. Unclick and then click &#34;Next step&#34; to bring up &#34;Seed Placement&#34; subheading.</li>
	<li>Click &#34;Validate Seeds&#34; and then click &#34;Next Step&#34; to bring up &#34;Neuron Reconstruction&#34; subheading.</li>
	<li>Under &#34;Neuron&#34;, click &#34;Interactive.&#34; Perform interactive tracing by following the direction from the program of the dendrites and right-clicking to complete the highlighted area traced by the program. After tracing all the dendrites, click &#34;Next Step.&#34;</li>
	<li>After a new window appears, save the new configuration with a desired title. Click &#34;Save.&#34;</li>
</ol>
9. Analysis
<ol>
	<li>Open the analysis program (see the table of materials).</li>
	<li>Click &#34;File&#34; from the top menu and then click &#34;Open Data File&#34;. Select the file of interest.</li>
	<li>Under the analysis subheading, select &#34;Sholl Analysis.&#34;</li>
	<li>In the &#34;Sholl Analysis&#34; window, set the starting radius to 10 &#181;m. Under &#34;Analysis,&#34; click the boxes &#34;Dendrites&#34; and &#34;Branch Orders.&#34;</li>
	<li>Right click the newly opened windows and click &#34;Excel Export.&#34; Click &#34;Save.&#34;</li>
	<li>Under the analysis, click &#34;Branched Structure Analyses.&#34; Select &#34;All Possible Analyses.&#34;</li>
	<li>Right click the newly opened window and click &#34;Excel Export&#34;24.</li>
</ol>

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The authors declare that they have no competing financial interests.

Compared to more modern techniques, the Golgi-Cox method has several advantages that make it the preferred method for examining spine morphology: 1) The staining can be used for essentially any tissue, 2) A basic light microscope setup is all that is needed to acquire Golgi-based images, 3) The Golgi-Cox imaging is faster than confocal imaging, and 4) Golgi stained sections are viable for several months to years longer than samples that are fluorescently-labeled. Even with these advantages, the Golgi-Cox method still has...

Dendrites are the largest portion of neurons that receive and process presynaptic input1. Their dendritic processes have a complex geometry, where the proximal branches have a larger diameter than the distal branches. As dendrites develop, they form several connections with other neurons in a process referred to as dendritic arborization. The extent and pattern of this branching determines the amount of synaptic inputs that a dendrite can adequately process2.
Dendritic arborization is a necessary process for activity-dependent plasticity and proper development of neuronal circuits. Extension, retraction, branching, and synaptogenesis are intricate processes that include intrinsic genetic programs and influences from extrinsic factors. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation3,4. The alterations in dendritic complexity are associated with pathophysiological and behavioral changes5. Abnormalities are related to several disease states, including Fragile X Syndrome and Down Syndrome6.
Dendritic spines are the specialized subcellular compartments of the dendritic arbors that receive excitatory input within the central nervous system. There are three morphological classes of dendritic spines, with the name of each class based on their size and shape: 1) mushroom spines, which have complex postsynaptic densities with more glutamate receptors than other spines7; 2) stubby spines, which lack a stem; and 3) thin spines, which consist of a protracted, narrow stem and a globular head8. The dendritic spine volume is used in part to define them, with thin spines generally smaller (0.01 &#181;m3) compared to mushroom spines (0.8 &#181;m3)9,10. The spines stabilize with maturation. For example, the thin spines either retract after a few days or develop into mushroom spines. Alternatively, the mushroom spines are relatively stable and can survive for an extended period.The strength of the neuronal connections is thought to be based on the number of spines and/or their volume11,12,13.
The classical Golgi staining method and its more modern variations have all been useful for examining the dendritic spine morphology and density. One unique aspect of the Golgi staining is that it randomly stains about 5% of the total neurons, which allows for the tracing of individual neurons14,15. Although the exact mechanism in which the Golgi method stains individual neurons is still unknown, the principle of the method is based on the crystallization of silver chromate (Ag2CrO4)16,17. There are three main types of the Golgi method: the rapid Golgi, the Golgi-Cox, and the Golgi-Kopsch18,19. All three methods start with an initial incubation phase in chromium salts for several days to months, but there are certain key differences between them. The rapid Golgi uses osmium tetroxide in the first step, whereas the Golgi-Kopsch includes paraformaldehyde. The staining in both the rapid-Golgi and the Golgi-Kopsch is followed by an incubation in a 1-2% silver nitrate solution for about 7 days. The Golgi-Cox method uses mercuric chloride and potassium dichromate instead of silver nitrate and has an impregnation time of 2-4 weeks. The tissues are then sectioned and quickly placed in a diluted ammonia solution, followed by a photographic fixer to remove salts. Of the three types, the Golgi-Cox method is thought to be the best at staining the dendritic arbors without much background interference, in part, because the crystal artifacts do not occur on the surface of the tissue (unlike in the rapid Golgi method)17,20,21.
The present method is a slight variation of the protocol provided with a commercial Golgi staining kit, and was designed to assess how a relatively low dose of the 5-Fu would affect the dendritic morphological characteristics and the spine density. Any data acquired could provide further insight into how chemotherapeutic treatment affects the neuronal circuitry.

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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.

The effects of the 5-Fu treatment on the dendritic arborization and complexity in the hippocampus of Golgi-stained brain sections were quantified and traced using a commercially available imaging software. After tracing, the dendritic arborization, the spine density, and the spine morphology were analyzed using Sholl analysis and the dendritic complexity index (DCI). Sholl analysis is a quantitative analytical method that can be used to determine the dendritic arbor morphology<sup class="...

Watch this Scientific Journal Video about Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method at JoVE.com

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

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 ...

assessment-hippocampal-dendritic-complexity-aged-mice-using-golgi-cox

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

14.9K Views.  University of Arkansas for Medical Sciences. 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.

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

Assessment of Motor Balance and Coordination in Mice using the Balance Beam

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice. Fine motor coordination and balance can be assessed by the beam walking assay. The goal of this test is for the mouse to stay upright and walk across an elevated narrow beam to a safe platform. This test takes place over 3 consecutive days: 2 days of training and 1 day of testing. Performance on the beam is quantified by measuring the time it takes for the mouse to traverse the beam and the number of paw slips that occur in the process. Here we report the protocol used in our laboratory, and representative results from a cohort of C57BL/6 mice. This task is particularly useful for detecting subtle deficits in motor skills and balance that may not be detected by other motor tests, such as the Rotarod.

Deficits in fine motor coordination can be assessed with the balance beam test. Performance on the beam is quantified by the speed at which the beam is traversed and the number of times the mouse slips on the beam.

Brain injury, genetic manipulations, and pharmacological treatments can result in alterations of motor skills in mice.  Fine motor coordination and ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6. Application of optogenetics7,8 in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17 or halorhodopsin18-20, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Here we describe a protocol for optogenetic manipulation of motoneuronal activity while monitoring changes in motor output (muscle contraction) in semi-intact Drosophila larvae using lasers within a conventional confocal microscope. This technique enables researchers to achieve local perturbation of neural activity in a few neuromeres to elucidate the dynamics of motor circuits.

Drosophila larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the ...

Visualizing the Effects of a Positive Early Experience, Tactile Stimulation, on Dendritic Morphology and Synaptic Connectivity with Golgi-Cox Staining

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile stimulation is a positive early experience that mimics maternal licking and grooming in the rat. Exposing rat pups to this positive experience can be completed easily and cost-effectively by using highly accessible materials such as a household duster. Using a cross-litter design, pups are either stroked or left undisturbed, for 15 min, three times per day throughout the perinatal period. To measure the neuroplastic changes related to this positive early experience, Golgi-Cox staining of brain tissue is utilized. Owing to the fact that Golgi-Cox impregnation stains a discrete number of neurons rather than all of the cells, staining of the rodent brain with Golgi-Cox solution permits the visualization of entire neuronal elements, including the cell body, dendrites, axons, and dendritic spines. The staining procedure is carried out over several days and requires that the researcher pay close attention to detail. However, once staining is completed, the entire brain has been impregnated and can be preserved indefinitely for ongoing analysis. Therefore, Golgi-Cox staining is a valuable resource for studying experience-dependent plasticity.

This paper describes the procedures for tactile stimulation of rat pups and subsequent Golgi-Cox staining of neuronal morphology. Tactile stimulation is a positive experience that is administered in the perinatal period by stroking pups with a household duster. Golgi-cox staining is a reliable procedure permitting the visualization of entire neurons.

To generate longer-term changes in behavior, experiences must be producing stable changes in neuronal morphology and synaptic connectivity. Tactile ...

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 ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

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.

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 ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

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

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Chapters in this video

Assessment of Motor Balance and Coordination in Mice using the Balance Beam

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion

Visualizing the Effects of a Positive Early Experience, Tactile Stimulation, on Dendritic Morphology and Synaptic Connectivity with Golgi-Cox Staining

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

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

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

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Fluorescence and Bioluminescence Imaging of Subcellular Ca²⁺ in Aged Hippocampal Neurons