This work was supported by the Intramural Programs of the National Institute of Neurological Disorders and Stroke (NINDS) and National Institute on Deafness and Other Communication Disorders (NIDCD), of the National Institutes of Health (NIH). We thank the NINDS EM facility and the NIDCD advanced imaging core (code # ZIC DC 000081-03) for assistance.

Care and handling of animals were in accordance with NIH Animal Care and Use Committee Guidelines. Postnatal day (P) 15-21 Sprague-Dawley rats, injected with 1-1.2% CTB bilaterally through the superior colliculus, were maintained on a 12:12-hr light:dark cycle.
1. Retinal Tissue Fixation
<ol>
	<li>Assemble the following materials and tools: A dissecting microscope, 2 forceps with very fine tips, scissors, cellulose filter paper, plastic pipette and a microscope slide.</li>
	<li>Anesthetize the rat in a closed chamber with 2.0 ml halothane (an inhalant anesthetic). Determine adequate anesthetization by these methods: lack of withdrawal of rear paw after toe pinch, or lack of blink reflex. Then decapitate immediately with a guillotine. Remove the eyes with a pair of iris scissors and place in a glass dish containing 4% paraformaldehyde in 0.1 M phosphate buffer (PB) at pH 7.4.</li>
	<li>Using the dissecting microscope, remove the cornea by cutting off the front of the eyeball. Remove the lens and vitreous from the inner retinal surface with forceps.</li>
	<li>Peel the sclera with the two forceps until the retina is isolated from the eyecup.</li>
	<li>Cut the retina immediately into 100 - 200 &#181;m-thick strips with a razor, and subject to pH-shift fixation.</li>
	<li>Fix retina strips in 4% paraformaldehyde in 0.1 M PB at pH 6.0 for 20 - 30 min and then in 4% paraformaldehyde plus 0.01% glutaraldehyde at pH 10.5 for 10 - 20 min at&#160;RT.</li>
	<li>After several washes in PB with 0.15 mM CaCl2 (pH 7.4 at 4&#176;C), cryoprotect the retinal strips with glycerol (60 min each in 10%, 20%, 30%, then O/N in 30%) in 0.1 M PB prior to freeze substitution.</li>
</ol>
2. Freeze-substitution
NOTE: This freeze-substitution method is modified from an earlier published protocol 19,20. Also, it is crucial that the instruments are very cold (wear gloves); otherwise, the tissue may thaw partially when touched with the instruments. All of these steps are done within the AFS chamber and the instruments are never allowed to move above the rim of the chamber. Similarly, proper cooling of all chemicals used in the AFS is necessary.
<ol>
	<li>Plunge-freeze the retinal strips in liquid propane at -184&#176;C in an EM cryopreservation instrument (CPC). Using a fine brush, place the samples on small pieces of double-stick tape attached to the metal stubs that go on the end of the plunging rods (wick off extra buffer using the brush).</li>
	<li>Plunge the rods into the liquid propane and keep there for about 5 sec, and then transfer to the automatic freeze-substitution instrument (AFS) using a small transport chamber that is filled with liquid nitrogen (&#39;LN2 cooled cryo transfer container&#39;).</li>
	<li>After placing the frozen sample and instruments (forceps and scalpel) into the AFS, cool the instruments in the chamber for several minutes before touching the tissue, or cool them by placing the tips of the instruments for a few seconds into the small transport chamber (filled with liquid nitrogen (&#39;LN2&#160;cooled cryo transfer container&#39;)).&#160;&#160;</li>
	<li>Once in the AFS, remove the sample and tape from the stub using a fine scalpel. Also, keep the nitrogen gas flow control, TF (TF is described as a &#39;regulator control for LN2 vaporiser&#39;) open during these procedures.</li>
	<li>Prior to placing the frozen tissue into the flat-embedding holders (i.e., before setting up the AFS), cut a thin circle from a clear plastic sheet and place it into the bottom of the holder so as to line the bottom of each well. This allows relatively easier removal of the polymerized specimen blocks when finished. 
	NOTE: Previously, we used an alternative method to the flat embedding holders, and employing double Reichert+gelatin capsules (see Petralia and Wenthold, 1999) 20.</li>
	<li>Use the following automatic sequence in the AFS (using instrument terminology): T1 = -90&#176;C for 32 hr, S1 = increase temperature by 4&#176;C/hr&#160;(11.3 hr), T2 = -45&#176;C for 50 hr, S2 = increase temperature by 5&#176;C/hr&#160;(9 hr), T3 = 0&#176;C for 40 hr (total = 142.3 hr). It may take 2 - 4 hr to place samples in the AFS (at -90&#176;C) prior to starting the timed sequence.</li>
	<li>Immerse the frozen sections in 1.5% uranyl acetate in methanol at -90&#176;C for &#62;32 hr in the AFS. Place two pieces of tissues (typically) in each of the seven wells in the flat-embedding aluminum holder (three fit into the AFS).
	<ol>
		<li>Place the tissue+tape into the uranyl acetate/methanol in the wells and remove the tape later, just prior to beginning embedding medium (such as Lowicryl HM20) infiltration, if it is too difficult to remove the tissue from the tape.</li>
	</ol>
	</li>
	<li>Then increase the temperature stepwise to -45&#176;C (+4&#176;C per hr; in the automatic sequence).</li>
	<li>Wash the samples three times in precooled methanol, by using a fine-tipped plastic pipette to remove the old solution from each flat-embedding holder, and then using another standard plastic pipette to add the precooled fresh methanol.</li>
	<li>Then, using the same method, infiltrate the samples progressively with low temperature embedding resins such as embedding medium (HM20/methanol at 1:1 and 2:1, each for 2 hr, followed by pure resin for 2 hr and then change the resins and keep O/N).</li>
	<li>Change the resin again the next day, adjusting the level of the resin to reach just to the top edge of the wells.</li>
	<li>Polymerize the samples (-45&#176;C to 0&#176;C in automatic sequence; +5&#176;C per hr) with UV light for 40 hr.</li>
	<li>Then remove the samples from the AFS. Typically, sample blocks still show some pinkness in color. Place the samples close to the fluorescent light in the chemical fume hood, at RT O/N or longer until they appear completely clear.</li>
</ol>
3. Postembedding EM Immunogold Immunocytochemistry
NOTE: Postembedding immunocytochemistry is performed as described 23,24,25.
<ol>
	<li>Cut 1 &#181;m sections with ultramicrotome, stain sections with 1% toluidine-blue, and examine them for section orientation; orient the tissue block to achieve the optimal transverse plane of sectioning.</li>
	<li>Cut 70 nm thick ultrathin sections with ultramicrotome and collect them on Formvar-carbon-coated nickel slot grids.</li>
	<li>Wash grids one time in distilled H2O followed by a Tris-buffered saline three times (TBS, 0.05 M Tris buffer, 0.7% NaCl, pH 7.6) wash.</li>
	<li>Incubate grids in 20 &#181;l drops of 5% BSA in TBS for 30 min , and then in 20 &#181;l drops of a mixture containing goat anti-CTB (1:3,000) and an antibody to one NMDAR subunit (anti-rabbit GluN2A 1:50, GluN2B 1:30), or an anti-rabbit GluA2/3 (1:30) in TBS-Triton (TBST, 0.01% Triton X-100, pH 7.6) with 2% BSA and 0.02 M NaN3 O/N at RT.</li>
	<li>Wash grids on three separate drops (20 &#181;l) of TBS (pH 7.6) for 10, 10, and 20 min, followed by TBS (pH 8.2) for 5 min.</li>
	<li>Incubate grids for 2 hr on drops (20 &#181;l) of a mixture containing donkey anti-rabbit IgG (1:20) coupled to 10 nm gold particles and donkey anti-goat IgG (1:20) coupled to 18 nm gold particles in TBST (pH 8.2) with 2% BSA and 0.02 M NaN3.</li>
	<li>Wash grids in 20 &#181;l drops of TBS (pH 7.6) for 5, 5, and 10 min, then in ultrapure water and then dry them.</li>
	<li>Counterstain grids with 5% uranyl acetate and 0.3% lead citrate in distilled H2O for 8 and 5 min under dark conditions, respectively.</li>
	<li>For triple-labeling experiments, incubate grids O/N&#160;at RT with 20 &#181;l drops of a mixture of anti-goat CTB (1:3,000), anti-mouse PSD-95 (1:100), and anti-rabbit GluN2A (1:50). Then incubate grids for 2 hr on 20 &#181;l drops of a mixture of IgGs coupled to 18, 10, and 5 nm gold particles in TBST (pH 8.2) with 2% BSA and 0.02 M NaN3. Keep the same procedures as double labeling for washing and counterstaining between and after antibody incubation.</li>
	<li>Controls were performed in which the secondary antibodies were applied alone.</li>
	<li>View grids on an EM and digitalize images. Process final figures in Adobe Photoshop 6.0 only for brightness and contrast if it is necessary24.</li>
</ol>
4. Quantification
<ol>
	<li>Manually select areas of the inner plexiform layer (IPL) without holes or cracks at 8,000X magnification, then randomly photomontage the full depth of IPL at 25,000X magnification.</li>
	<li>Identify RGC dendrites at cone bipolar dyads when they a) contain the retrogradely transported CTB signal (large gold particles), b) exhibit well-defined membranes, clefts, and postsynaptic densities, c) contain at least two small gold particles within the PSD, or more than one small gold particle along the extrasynaptic plasma membrane.</li>
	<li>Collect 45 - 55 RGC dendrites, based on above criteria, for quantitative analysis.</li>
	<li>Measure the lengths of the PSD and the extrasynaptic plasma membrane of individual RGC dendrites with NIH ImageJ software. Count gold particles within 10 nm of the membrane as membrane-associated, based on an average thickness of 7 - 9 nm for the plasma membrane&#160;26.</li>
	<li>Calculate particle density as the number of gold particles per linear micrometer (gold/&#181;m). Measure the distance between the center of each gold particle and the middle of the PSD (for synaptic location) or the edge of the PSD (for extrasynaptic location).</li>
	<li>Perform statistical analysis by software. Use two-tailed t-tests to compare means. Conclude significance when P &#60;0.05. Unless indicated otherwise, report values as mean &#177; SEM18.</li>
</ol>

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The authors have no disclosures.

We have described four techniques for successful quantitative post-embedding immunogold EM: 1) short and weak fixation, 2) freeze-substitution, 3) post-embedding immunogold staining, and 4) quantification.
EM immunogold allows the detection of specific proteins in ultrathin tissue sections. Antibodies labeled with gold particles can be directly visualized using EM. While powerful in detecting the subsynaptic localization of a membrane receptor, EM immunogold can be technically challenging, and...

Glutamate is the major excitatory neurotransmitter in the retina1. Retinal ganglion cells (RGCs), receiving glutamatergic synaptic input from bipolar cells2, are the output neurons of the retina that send visual information to the brain. Physiological studies showed that synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs) 3,4,5. Although excitatory postsynaptic currents (EPSCs) in RGCs are mediated by AMPARs and NMDARs3,5,6,7,8 , spontaneous miniature EPSCs (mEPSCs) on RGCs exhibit only an AMPARs-mediated component 4,5,9. However, reducing glutamate uptake revealed an NMDAR component in spontaneous EPSCs5, suggesting that NMDARs on RGC dendrites may be located outside of excitatory synapses . Membrane-associated guanylate kinases (MAGUKs) such as PSD-95 that cluster neurotransmitter receptors, including glutamate receptors and ion channels at synaptic sites, also exhibit distinct subsynaptic expression patterns 10,11,12,13,14.
Over recent decades, confocal immunohistochemistry and pre-embedding electron microscope (EM) immunohistochemistry have been employed to study membrane receptor expression. Although confocal immunostaining reveals broad patterns of receptor expression, its lower resolution makes it impossible to use to distinguish subcellular location. Pre-embedding EM studies in mammalian retina indicate that NMDAR subunits are present in postsynaptic elements at cone bipolar cell ribbon synapses 15,16,17. This is in apparent contrast to physiological evidence. However, diffusion of reaction product is a well-known artifact in the pre-embedding immunoperoxidase method. Hence, this approach does not usually give statistically reliable data and may exclude distinction between localization to synaptic membrane versus extrasynaptic membrane 18,19,20,21. On the other hand, physiological and anatomical data are consistent with a synaptic localization of AMPARs on RGCs 3,5,7,9,22. Thus, glutamate receptors and MAGUKs at retinal ribbon synapse are localized not only to the postsynaptic but also to the perisynaptic or extrasynaptic membrane compartments. However, a high-resolution quantitative analysis of these membrane proteins in a retinal ribbon synapse is still needed.
Here, we developed a postembedding EM immunogold technique to examine the subsynaptic localization of NMDAR subunits, AMPAR subunits and PSD-95 followed by estimating the number, density and variability of these proteins at synapses onto rat RGCs labeled using cholera toxin subunit B (CTB) retrograde tracing methods.

<table><tbody><tr><td>Paraformaldehyde</td><td>EMS</td><td>15710</td><td></td></tr><tr><td>Glutaraldehyde</td><td>EMS</td><td>16019</td><td></td></tr><tr><td>NaH2PO4</td><td>Sigma</td><td>S9638</td><td></td></tr><tr><td>Na2HPO4</td><td>Sigma</td><td>7782-85-6</td><td></td></tr><tr><td>CaCl2</td><td>Sigma</td><td>C-8106</td><td></td></tr><tr><td>BSA</td><td>Sigma</td><td>A-7030</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>T-8787</td><td></td></tr><tr><td>NaOH</td><td>Sigma</td><td>221465</td><td></td></tr><tr><td>NaN3</td><td>JT Baker</td><td>V015-05</td><td></td></tr><tr><td>Glycerol</td><td>Gibco BRL</td><td>15514-011</td><td></td></tr><tr><td>Lowicryl HM 20</td><td>Polysciences</td><td>15924-1</td><td></td></tr><tr><td>Tris-Base</td><td>Fisher</td><td>BP151-500</td><td></td></tr><tr><td>Tris</td><td>Fisher</td><td>04997-100</td><td></td></tr><tr><td>Anti-GluN2A</td><td>Millipore</td><td>AB1555P</td><td>Dilution 1/50</td></tr><tr><td>Anti-GluN2B</td><td>Millipore</td><td>AB1557P</td><td>Dilution 1/30</td></tr><tr><td>Anti-GluA2/3</td><td>Millipore</td><td>AB1506</td><td>Dilution 1/30</td></tr><tr><td>Anti-PSD-95</td><td>Millipore</td><td>MA1&amp;ndash;046</td><td>Dilution 1/100</td></tr><tr><td>Donkey anti-rabbit IgG-10 nm gold particles</td><td>EMS</td><td>25704</td><td>Dilution 1/20</td></tr><tr><td>Donkey anti-mouse IgG-10 nm gold particles</td><td>EMS</td><td>25814</td><td>Dilution 1/20</td></tr><tr><td>Donkey anti-mouse IgG-5 nm gold particles</td><td>EMS</td><td>25812</td><td>Dilution 1/20</td></tr><tr><td>Donkey anti-goat IgG-18 nm gold particles</td><td>Jackson ImmunoResearch</td><td>705-215-147</td><td>Dilution 1/20</td></tr><tr><td>Formvar-Carbon coated nickel-slot grids.</td><td>EMS</td><td>FCF2010-Ni</td><td></td></tr><tr><td>Uranyl acetate</td><td>EMS</td><td>22400-1</td><td></td></tr><tr><td>Methanol</td><td>EMS</td><td>67-56-1</td><td></td></tr><tr><td>Lead citrate</td><td>Leica</td><td></td><td></td></tr><tr><td>Leica EM AFS</td><td>Leica</td><td></td><td></td></tr><tr><td>Leica EM CPC</td><td>Leica</td><td></td><td></td></tr><tr><td>Ultramicrotome</td><td>Leica</td><td></td><td></td></tr><tr><td>JEOL 1200 EM</td><td>JEOL</td><td></td><td></td></tr><tr><td>liquid nitrogen&amp;nbsp;</td><td>Roberts Oxygen</td><td></td><td></td></tr><tr><td>Propane</td><td>Roberts Oxygen</td><td></td><td></td></tr><tr><td>CTB</td><td>List Biological Laboratories</td><td>104</td><td>1 - 1.2%</td></tr><tr><td>Anti-CTB</td><td>List Biological Laboratories</td><td>703</td><td>Dilution 1/4,000</td></tr></tbody></table>

high-resolution quantitative immunogold analysis of membrane receptors at retinal ribbon synapses

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA receptors (NMDARs) and AMPA receptors (AMPARs). Physiological data have indicated that glutamate receptors at RGCs are expressed not only in postsynaptic but also in perisynaptic or extrasynaptic membrane compartments. However, precise anatomical locations for glutamate receptors at RGC synapses have not been determined. Although a high-resolution quantitative analysis of glutamate receptors at central synapses is widely employed, this approach has had only limited success in the retina. We developed a postembedding immunogold method for analysis of membrane receptors, making it possible to estimate the number, density and variability of these receptors at retinal ribbon synapses. Here we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of retinal fixation, 2) freeze-substitution, 3) postembedding immunogold electron microscope (EM) immunocytochemistry and, 4) quantitative visualization of glutamate receptors at ribbon synapses.

The results presented here demonstrate strikingly different subsynaptic localization patterns of GluA 2/3 and NMDARs on RGC dendrites in rat retina, as described previously 24,25. 77% of GluA 2/3 immunogold particles in RGC dendritic profiles were located within the PSD (Figure 1A), similar to most central synapses. However, NMDARs were located either synaptically or extrasynaptically. 83% of GluN2A immunogold particles were localized in the PSD (Figure...

Watch this Scientific Journal Video about High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses at JoVE.com

High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Retinal ganglion cells (RGCs) receive excitatory glutamatergic input from bipolar cells. Synaptic excitation of RGCs is mediated postsynaptically by NMDA ...

high-resolution-quantitative-immunogold-analysis-membrane-receptors

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

7.5K Views.  National Institutes of Health. The postembedding immunogold method is one of the most effective ways to provide high-resolution analyses of the subcellular localization of specific molecules. Here we describe a protocol to quantitatively analyze glutamate receptors at retinal ribbon synapses.

Video: High-Resolution Quantitative Immunogold Analysis of Membrane Receptors at Retinal Ribbon Synapses

Quantifying Synapses: an Immunocytochemistry-based Assay to Quantify Synapse Number

One of the most important goals in neuroscience is to understand the molecular cues that instruct early stages of synapse formation. As such it has become imperative to develop objective approaches to quantify changes in synaptic connectivity. Starting from sample fixation, this protocol details how to quantify synapse number both in dissociated neuronal culture and in brain sections using immunocytochemistry. Using compartment-specific antibodies, we label presynaptic terminals as well as sites of postsynaptic specialization. We define synapses as points of colocalization between the signals generated by these markers. The number of these colocalizations is quantified using a plug in Puncta Analyzer (written by Bary Wark, available upon request, c.eroglu@cellbio.duke.edu) under the ImageJ analysis software platform. The synapse assay described in this protocol can be applied to any neural tissue or culture preparation for which you have selective pre- and postsynaptic markers. This synapse assay is a valuable tool that can be widely utilized in the study of synaptic development.

This protocol details how to quantify synapse number both in dissociated neuronal culture and in brain sections using immunocytochemistry. Using compartment-specific antibodies, we label presynaptic terminals as well as sites of postsynaptic specialization. We define synapses as points of colocalization between the signals generated by these markers.

One of the most important goals in neuroscience is to understand the molecular cues that instruct early stages of synapse formation. As such it has become ...

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

Biology

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 ...

Immunohistochemical and Calcium Imaging Methods in Wholemount Rat Retina

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for immunohistochemistry and, 2) calcium imaging for the study of voltage gated calcium channel (VGCC) mediated calcium signaling in retinal ganglion cells. The calcium imaging method we describe circumvents issues concerning non-specific loading of displaced amacrine cells in the ganglion cell layer.

Immunohistochemistry protocols are used to study the localization of a specific protein in the retina. Calcium imaging techniques are employed to study calcium dynamics in retinal ganglion cells and their axons.

In this paper we describe the tools, reagents, and the practical steps that are needed for: 1) successful preparation of wholemount retinas for ...

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

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

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

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

Imaging the Human Immunological Synapse

The purpose of the method is to generate an immunological synapse (IS), an example of cell-to-cell conjugation formed by an antigen-presenting cell (APC) and an effector helper T lymphocyte (Th) cell, and to record the images corresponding to the first stages of the IS formation and the subsequent trafficking events (occurring both in the APC and in the Th cell). These events will eventually lead to polarized secretion at the IS. In this protocol, Jurkat cells challenged with Staphylococcus enterotoxin E (SEE)-pulsed Raji cells as a cell synapse model was used, because of the closeness of this experimental system to the biological reality (Th cell-APC synaptic conjugates). The approach presented here involves cell-to-cell conjugation, time-lapse acquisition, wide-field fluorescence microscopy (WFFM) followed by image processing (post-acquisition deconvolution). This improves the signal-to-noise ratio (SNR) of the images, enhances the temporal resolution, allows the synchronized acquisition of several fluorochromes in emerging synaptic conjugates and decreases fluorescence bleaching. In addition, the protocol is well matched with the end point cell fixation protocols (paraformaldehyde, acetone or methanol), which would allow further immunofluorescence staining and analyses. This protocol is also compatible with laser scanning confocal microscopy (LSCM) and other state-of-the-art microscopy techniques. As a main caveat, only those T cell-APC boundaries (called IS interfaces) that were at the right 90&#176; angle to the focus plane along the Z-axis could be properly imaged and analyzed. Other experimental models exist that simplify imaging in the Z dimension and the following image analyses, but these approaches do not emulate the complex, irregular surface of an APC, and may promote non-physiological interactions in the IS. Thus, the experimental approach used here is suitable to reproduce and to confront some biological complexities occurring at the IS.

This protocol images both the immunological synapse formation and the subsequent polarized secretory traffic towards the immunological synapse. Cellular conjugates were formed between a superantigen-pulsed Raji cell (acting as an antigen-presenting cell) and a Jurkat clone (acting as an effector helper T lymphocyte).

The purpose of the method is to generate an immunological synapse (IS), an example of cell-to-cell conjugation formed by an antigen-presenting cell (APC) ...

Immunology and Infection

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Optimized Protocol for Retinal Organoid Sample Preparation for TEM

Preparing Retinal Organoid Samples for Transmission Electron Microscopy

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem cells (iPSCs) under specific conditions. Synapse development and maturation in ROs have been studied immunocytochemically and functionally. However, the direct evidence of the synaptic contact ultrastructure is limited, containing both special ribbon synapses and conventional chemical synapses. Transmission electron microscopy (TEM) is characterized by high resolution and a respectable history elucidating retinal development and synapse maturation in humans and various species. It is a powerful tool to explore synaptic structure in ROs and is widely used in the research field of ROs. Therefore, to better explore the structure of RO synaptic contacts at the nanoscale and obtain high-quality microscopic evidence, we developed a simple and repeatable method of RO TEM sample preparation. This paper describes the protocol, reagents used, and detailed steps, including RO fixation preparation, post fixation, embedding, and visualization.

Author Spotlight: Analyzing the Synaptic Ultrastructure in Mature Retinal Organoids Using TEM

This protocol provides an optimized and elaborate preparation procedure for retinal organoid samples for transmission electron microscopy. It is suitable for applications that involve the analysis of synapses in mature retinal organoids.

Retinal organoids (ROs) are a three-dimensional culture system mimicking human retinal features that have differentiated from induced pluripotent stem ...

Synaptic Circuits Analysis and Protein Localization in the Mouse Retina

Investigating Retinal Circuits and Molecular Localization by Pre&#45;Embedding Immunoelectron Microscopy

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is characterized by unique features and distinct neurotransmitters, determining its role and functional significance. Given the intricate cell types within its structure, the complexity of neuronal circuits in the retina poses challenges for exploration. To better investigate retinal circuits and cross-talk, such as the link between cone and rod pathways, and precise molecular localization (neurotransmitters or neuropeptides), such as the presence of substance P-like immunoreactivity in the mouse retina, we employed a pre-embedding immunoelectron microscopy (immuno-EM) method to explore synaptic connections and organization. This approach enables us to pinpoint specific intercellular synaptic connections and precise molecular localization and could play a guiding role in exploring its function. This article describes the protocol, reagents used, and detailed steps, including (1) retina fixation preparation, (2) pre-embedding immunostaining, and (3) post-fixation and embedding.

Author Spotlight: Understanding the Ultrastructural Basis of Retinal Synaptic Connectivity and Neurotransmitter Localization in Mice 

This protocol outlines the detailed steps of pre-embedding immunoelectron microscopy, with a focus on exploring synaptic circuits and protein localization in the retina.

The retina comprises numerous cells forming diverse neuronal circuits, which constitute the first stage of the visual pathway. Each circuit is ...

Immunolabeling of Neuronal Membrane Proteins in a Freeze-fractured Specimen of Mouse Brain Tissue

<a href='https://appjove.unicartagenaproxy.elogim.com/v/53853/combined-optogenetic-freeze-fracture-replica-immunolabeling-to'>Source: Sch&#246;nherr, S., et. al. Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate Receptors in the Mouse Amygdala. J. Vis. Exp. (2016)</a>This video demonstrates the immunolabelling of freeze-fractured replicas of mouse brain tissue for electron microscopy. The fractured and coated replicas are digested to expose receptors. Primary and gold-tagged secondary antibodies are added to label specific neuronal membrane receptors. The replica is mounted on a grid and visualized under an electron microscope.

Source: Sch&#246;nherr, S., et. al. Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate ...