This work was supported by Sacred Heart University Undergraduate Research InitiativeGrants.

All experimental procedures are approved by the Sacred Heart University Institutional Animal Care and Use Committee and are in accordance with the NIH Guide for the Care and Use of Animals.
1. Isolation and infiltration of brain tissue
<ol>
	<li>Premix solutions A and B of the Golgi staining kit 24 h prior to use and keep in dark bottles and/or in dark. Make approximately 80 mL of solution A and B mix which is sufficient to change the solution after 24 h. Store in airtight bottles. 
	NOTE: Perfusion with either saline or paraformaldehyde is not necessary.</li>
	<li>Sacrifice rats by guillotine following carbon dioxide euthanasia and remove brains within seconds. Rinse brains in saline if required but is not necessary.
	<ol>
		<li>Place the brain, cortex down so that the hypothalamus is visible because the cuts are made anterior and posterior to it, on a non-porous surface. Cut into an anterior (contains the prefrontal cortex) and posterior (contains the hippocampus) block as shown in Figure 1.</li>
	</ol>
	</li>
	<li>Place blocks into the premixed solutions of A and B, making sure that they are well immersed in the solution. Ensure that the volume of solutions A and B is sufficient to immerse the blocks. Store blocks in either brown bottles or clear bottles covered with foil (to keep light out) in the dark at room temperature.</li>
	<li>Replace solution after 24 h and keep in the dark for another 13 days at room temperature. 
	NOTE: If used with mouse brains, there is no need to block, and the entire brain can be placed in the solution. If staining brain areas, which are different in size, the time in solutions A and B may have to be determined by trial and error.</li>
	<li>After two weeks the tissue is infiltrated well with solutions A and B, transfer the blocks to the cryoprotectant solution (solution C in the Golgi staining kit), and leave them for 48-72 h at 4 &#176;C. 
	NOTE: After cryoprotection, blocks may be frozen until further processing. Also, note that all solutions from the kit are collected and disposed of as hazardous waste.</li>
	<li>Freeze brain, cortex down, on a glass slide on dry ice. Once frozen either cut on the cryostat or store at -80 &#176;C until sectioning using a cryostat. 
	&#8203;NOTE: Do not freeze in liquid nitrogen as this produces cracks in the tissue.</li>
</ol>
2. Sectioning of brain tissues
<ol>
	<li>Place a small amount of tissue medium on a pre-cooled cryostat chuck. Mount blocks on cryostat chucks by thawing one side of the block slightly in hand (gloved) and placing it on the tissue medium. 
	NOTE: It is not necessary to embed the tissue. The block containing the hippocampus is somewhat easier to cut than the block with the prefrontal cortex because the latter is anterior to the corpus callosum and the two halves are separate.</li>
	<li>Cut 100 &#181;m sections on a cryostat at -22 &#176;C and mount onto subbed slides. Sections up to 150 &#181;m of thickness can be used. Try to mount 3-4 coronal sections per slide as this decreases the number of slides that require processing. Use one of the following techniques to keep the sections frozen till they get on the slide. 
	NOTE: These sections are not fixed in the usual way and therefore they tend to melt quickly. Allow the sections to melt on the slide. If they begin to melt before being placed on the slide, it is impossible to get them to be flat on the slide. There are several techniques to do this.
	<ol>
		<li>Use a freezing spray on the knife and the block. Depending on the temperature and humidity in the room this may be necessary for every section. Use either the antiroll plate or a brush to keep the section flat while cutting. 
		NOTE: Make sure to have enough freezing spray. Several cans can be used when cutting 20 blocks.</li>
		<li>Thaw mount, if possible, by using a room temperature slide and quickly appose it to the section. With practice, one can mount several sections at once. If this doesn&#39;t work, keep slides in the cryostat so they are very cold and transfer the section with cold forceps or a cold paintbrush and then thaw mount.</li>
	</ol>
	</li>
	<li>Clean the knife with paper wipes between slices. If the knife requires more cleaning, use 100% ethanol (EtOH), and let it dry before cutting the next slice.</li>
	<li>Once mounted onto the slide, place the slide flat on cardboard slide trays and allow sections to dry at room temperature (for several hours to a maximum of 48 h) in the dark. Do not cover the slide tray. Ensure that the sections are completely dry before Golgi impregnation. Store flat, in the dark, on slide trays until Golgi impregnation. 
	&#8203;NOTE: Drying the sections does not cause any damage.</li>
</ol>
3. Staining and dehydration of brain tissue
<ol>
	<li>Perform staining in glass staining dishes. Mix solutions D and E immediately prior to staining. As per the protocol, add one part of solution D, one part of solution E, and two parts of distilled water. For glass staining dishes, make a 200 mL solution to completely submerge the sections. For Koplin jars, this requires less volume.</li>
	<li>Place slides in racks spaced far enough apart to allow solution access to sections.</li>
	<li>Place sections in distilled water for 4 min (2x) before placing them into the Golgi impregnation staining solution for 10 min. Change the staining solution every 70 sections. Change the distilled water when it turns yellow. 
	NOTE: The timing for the staining step is critical. Too short does not allow enough staining and too long causes over staining, making the dendrites hard to separate when doing analysis. Once out of the staining solution the timing is less critical.</li>
	<li>Dehydrate the sections as follows: 70% EtOH (5 min), 95% EtOH (5 min; 2x), 100% EtOH (5 min; 2x), clearing agent (5 min; 3x). Change all solutions often, especially the 100% EtOH and clearing agent to ensure the sections remain anhydrous. 
	NOTE: It is not necessary to go through a 50% EtOH step. Counterstaining is not needed for cell visualization because the Golgi impregnation provides sufficient contrast. The use of other clearing agents has not worked as well as the one used here (see Table of materials).</li>
	<li>Coverslip with glass coverslips that are 60 mm long with a generous amount of mounting medium. Make sure that there are, as few air bubbles as possible. If needed, carefully remove, and redo the coverslip (can be done even weeks later). Do this very slowly in order not to damage the section (can be done because of the large amount of mounting medium). 
	NOTE: A large amount of mounting medium is somewhat messy but still necessary because enough mounting medium must be used to cover the thick 100 &#181;m sections.</li>
	<li>To ensure that only one coverslip is placed on the sections, separate the coverslips in advance of the process.</li>
	<li>Once cover slipped, dry slides flat on any non-porous paper for 3-5 days, moving them slightly, especially after the first day, to avoid sticking. After 3-5 days transfer the slides to slide holders and, ideally, dry slides for at least 3 weeks before examining them. Keep slides flat to decrease the possibility of air bubbles forming.</li>
</ol>
4. Determination of dendritic spine density
<ol>
	<li>For analysis of dendritic spine density in pyramidal neurons of both the mPFC and the CA1 region of the hippocampus, examine the most lateral secondary basal dendrites and the most lateral tertiary apical dendrites as described in step 4.1.1 (Figure 2).
	<ol>
		<li>Choose a dendrite, measure the length of the dendrite using an image analysis program, count the spines on the dendrites using a hand counter, and record both length and number of spines.</li>
	</ol>
	</li>
	<li>Study and analyze six cells per region (mPFC, CA1) per brain. Quantify a minimum of six brains per group as previously described7,8. Choose neurons that meet the following criteria for analysis: cell bodies and dendrites are well impregnated; dendrites are distinguishable from adjacent cells and are continuous.</li>
	<li>Count the spines at 1000x (oil-immersion) by hand counting with a light microscope and measure dendritic length using an image analysis program. Calculate spine density by dividing the spine number by the length of the dendrite and express data as the number of spines/ 10 &#181;m dendrite. 
	NOTE: There are far more sophisticated methods for differentiating spine subtypes and dendritic architecture that can be used, but hand counting with a light microscope at 1000x can give a rapid result that can then determine if further investigation is necessary. Although every effort is made to consistently sample similar dendrites, there are variations in thickness that may affect the counting.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Environmental+enrichment+prevents+acute+restraint+stress-induced+anxiety-related+behavior+but+not+changes+in+basolateral+amygdala+spine+density.">Environmental enrichment prevents acute restraint stress-induced anxiety-related behavior but not changes in basolateral amygdala spine density.</a> Psychoneuroendocrinology. 98, 6-10 (2018).</li><li>Trzesniewski, J., Altmann, S., J&#228;ger, L., Kapfhammer, J. 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The authors have nothing to disclose.

The present protocol describes a method of Golgi impregnation that allows for rapid simultaneous processing of many sections. It is an improvement over previously described5 more labor-intensive methods and consistently yields impregnated neurons for analysis. In addition, there is less exposure to toxic chemicals used in Golgi impregnation. The most challenging part of the process is getting the sections to be flat on the slides, which takes considerable practice. Keeping everything as cold as po...

The method of using potassium dichromate and silver nitrate to label neurons was first described by Camillo Golgi1,2 and subsequently used by Santiago Ramon y Cajal to produce an immense body of work differentiating neuronal and glial subtypes. A recently published book with his illustrations is now available3. Following Ramon y Cajal&#39;s studies, which were published more than 100 years ago, very little Golgi impregnation was used. Golgi impregnation is a laborious process that allows three-dimensional visualization of neurons with a light microscope. There have been numerous modifications of the Golgi method over the years to make the method easier and the staining more consistent4. In 1984, Gabbott and Somogyi5 described the single section Golgi impregnation procedure which allowed for more rapid processing. This Golgi impregnation method requires perfusion with 4% paraformaldehyde and 1.5% picric acid, post-fixation followed by vibratome sectioning into a bath of 3% potassium dichromate. Sections are mounted onto glass slides, the four corners of coverslips glued so that when immersed in silver nitrate, diffusion is gradual. Coverslips are then popped off, sections are dehydrated, and eventually cover slipped permanently with mounting medium. This technique was successfully used to label neurons and glia6,7,8 in the hippocampus. The rapid Golgi method described here is an improvement because there is far less exposure to both potassium dichromate and silver nitrate and no paraformaldehyde and picric acid are used. In addition, although cells that were impregnated using modifications of the Gabbott and Somogyi5 method could be analyzed, often the sections were over-or under-exposed or fell off the slides during the dehydration step and generally, several experiments had to be pooled to have enough cells for analysis.
The present protocol describes the use of the Golgi staining kit (see Table of materials) to label dendrites and dendritic spines in the hippocampus and medial prefrontal cortex (mPFC) of the rat. The advantages of this method over previous ones are that it is rapid, there is less exposure to noxious chemicals for the researcher and there is consistent staining of neurons. The protocol described below has been used with minor modifications to assess dendritic spine density in the hippocampus and mPFC of the rat in many studies9,10,11,12,13,14,15.

<table><tbody><tr><td>Cardboard slides trays</td><td>Fisher Scientific</td><td>12-587-10</td><td></td></tr><tr><td>Coverslips 24 x 60mm</td><td>Fisher Scientific</td><td>12-545-M</td><td></td></tr><tr><td>FD Rapid GolgiStain kit</td><td>FD Neurotechnologies</td><td>PK 401</td><td>Stable at RT in the dark for months; Golgi staining kit</td></tr><tr><td>Freezing Spray</td><td>Fisher Scientific</td><td>23-022524</td><td></td></tr><tr><td>HISTO-CLEAR</td><td>Fisher Scientific</td><td>50-899-90147</td><td>clearing agent</td></tr><tr><td>NCSS Software</td><td>Kaysville, UT, USA</td><td></td><td></td></tr><tr><td>Permount</td><td>Fisher Scientific</td><td>SP-15-100</td><td>mounting medium</td></tr><tr><td>Superfrost Plus Microscope slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Tissue Tek CTYO OCT Compound</td><td>Fisher Scientific</td><td>14-373-65</td><td>Used to mount brains on cryostat chuck</td></tr></tbody></table>

rapid golgi stain for dendritic spine visualization in hippocampus and prefrontal cortex

Golgi impregnation, using the Golgi staining kit with minor adaptations, is used to impregnate dendritic spines in the rat hippocampus and medial prefrontal cortex. This technique is a marked improvement over previous methods of Golgi impregnation because the premixed chemicals are safer to use, neurons are consistently well impregnated, there is far less background debris, and for a given region, there are extremely small deviations in spine density between experiments. Moreover, brains can be accumulated after a certain point and kept frozen until further processing. Using this method any brain region of interest can be studied. Once stained and cover slipped, dendritic spine density is determined by counting the number of spines for a length of dendrite and expressed as spine density per 10 &#181;m dendrite.

Using the rapid Golgi method, cells are consistently well impregnated so that there are plenty of cells to analyze. This is a marked improvement over prior methods where experiments had to be pooled to have enough data for analysis. Therefore, more samples can be processed at once and brains can be stored frozen until processing. Examples of Golgi impregnated cells in the CA1 region of the hippocampus are shown at low and high power in Figure 3. Counting of spines in a given region yields co...

Watch this Scientific Journal Video about Rapid Golgi Stain for Dendritic Spine Visualization in Hippocampus and Prefrontal Cortex at JoVE.com

Rapid Golgi Stain for Dendritic Spine Visualization in Hippocampus and Prefrontal Cortex

The protocol describes a modification of the rapid Golgi method, which can be adapted to any part of the nervous system, for staining neurons in the hippocampus and medial prefrontal cortex of the rat.

Golgi impregnation, using the Golgi staining kit with minor adaptations, is used to impregnate dendritic spines in the rat hippocampus and medial ...

rapid-golgi-stain-for-dendritic-spine-visualization-hippocampus

Universidad de Cartagena UDEC

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Neuroscience

6.1K Views.  Donald and Barbara Zucker School of Medicine at Hofstra/Northwell. The protocol describes a modification of the rapid Golgi method, which can be adapted to any part of the nervous system, for staining neurons in the hippocampus and medial prefrontal cortex of the rat.

Video: Rapid Golgi Stain for Dendritic Spine Visualization in Hippocampus and Prefrontal Cortex

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Analysis of Dendritic Spine Morphology in Cultured CNS Neurons

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These structures are rich in actin and have been shown to be highly dynamic. In response to classical Hebbian plasticity 
as well as neuromodulatory signals, dendritic spines can change shape and number, which is thought to be critical for the refinement of neural 
circuits and the processing and storage of information within the brain. Within dendritic spines, a complex network of proteins link extracellular 
signals with the actin cyctoskeleton allowing for control of dendritic spine morphology and number. Neuropathological studies have demonstrated that 
a number of disease states, ranging from schizophrenia to autism spectrum disorders, display abnormal dendritic spine morphology or numbers. 
Moreover, recent genetic studies have identified mutations in numerous genes that encode synaptic proteins, leading to suggestions that these 
proteins may contribute to aberrant spine plasticity that, in part, underlie the pathophysiology of these disorders. In order to study the potential 
role of these proteins in controlling dendritic spine morphologies/number, the use of cultured cortical neurons offers several advantages. Firstly, 
this system allows for high-resolution imaging of dendritic spines in fixed cells as well as time-lapse imaging of live cells. Secondly, this in 
vitro system allows for easy manipulation of protein function by expression of mutant proteins, knockdown by shRNA constructs, or pharmacological 
treatments. These techniques allow researchers to begin to dissect the role of disease-associated proteins and to predict how mutations of these 
proteins may function in vivo.

Numerous recent studies have identified mutations in synaptic proteins associated with brain pathologies. Primary cultured cortical neurons offer great flexibility in examining the effects of these disease-associated proteins on dendritic spine morphology and motility.

Dendritic spines are the sites of the majority of excitatory connections within the brain, and form the post-synaptic 
compartment of synapses. These ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

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 ...

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 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 ...

Imaging Neurons within Thick Brain Sections Using the Golgi-Cox Method

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within histological brain samples. While it is preferable from a practical perspective to prepare brain sections at the greatest thickness possible, in order to increase the probability of identifying stained neurons that are fully contained within single sections, this approach is limited from a technical perspective by the working distance of high-magnification microscope objectives. We report here a protocol to stain neurons using the Golgi-Cox method in mouse brain sections that are cut at 500 &#956;m thickness, and to visualize neurons throughout the depth of these sections using an upright microscope fitted with a high-resolution 30X 1.05 N.A. silicone oil-immersion objective that has an 800 &#956;m working distance. We also report two useful variants of this protocol that may be employed to counterstain the surface of mounted brain sections with the cresyl violet Nissl stain, or to freeze whole brains for long-term storage prior to sectioning and final processing. The main protocol and its two variants produce stained thick brain sections, throughout which full neuron dendritic&#160;trees and dendrite spines may be reliably visualized and quantified.

We present a protocol for using the Golgi-Cox staining method in thick brain sections, in order to visualize neurons with long dendritic&#160;trees contained within single tissue samples. Two variants of this protocol are also presented that involve cresyl violet counterstaining, and the freezing of unprocessed brains for long-term storage.

The Golgi-Cox method of neuron staining has been employed for more than two hundred years to advance our understanding of neuron morphology within ...

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 ...

Serial Block-Face Scanning Electron Microscopy (SBEM) for the Study of Dendritic Spines

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In addition, some features of dendritic spines, such as volume of the spine and post-synaptic density (PSD) (representing post-synaptic part of the synapse), presence of presynaptic terminal, and smooth endoplasmic reticulum or atypical form of PSD (e.g., multi-innervated spines), can be observed only with 3D EM. By employing serial block-face scanning electron microscopy (SBEM) it is possible to obtain 3D EM data easier and in a more reproducible manner than when performing traditional serial sectioning. Here we show how to prepare mouse hippocampal samples for SBEM analysis and how this protocol can be combined with immunofluorescence study of dendritic spines. Mild fixation perfusion allows us to perform immunofluorescence studies with light microscopy on one half of the brain, while the other half was prepared for SBEM. This approach reduces the number of animals to be used for the study.

Serial Block-Face Scanning Electron Microscopy SBEM for the Study of Dendritic Spines

Serial Block-Face Scanning Electron Microscopy (SBEM) is applied to image and analyze dendritic spines in the murine hippocampus.

Three-dimensional electron microscopy (3D EM) gives a possibility to analyze morphological parameters of dendritic spines with nanoscale resolution. In ...