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

Summary

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.

Abstract

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 ultrastructure^3-5.

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 function^8,9 . 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.

Protocol

1. Fixation

Fixatives are carcinogenic; wear gloves and handle the fixatives in a fume hood. Unless otherwise noted, all incubations are done on ice and all solutions should be filtered before use. Use electron microscopy-grade reagents.

Day 1

1. After experimental conditions (e.g. viral injection, drug treatment), place the membrane with organotypic hippocampal slices in a 60 x 15 mm polystyrene Petri dish containing ice-cold 0.1 M phosphate buffer (Sorensen's phosphate buffer), pH 7.3. Add 1 - 2 ml of 0.1 M phosphate buffer directly on top of the slices to keep them cold.

CRITICAL STEP: Always use freshly prepared buffers and fixatives. Make sure the pH of buffer is within desired range. A failure to do so could damage cellular structures.

2. To isolate the CA1 subfield of the slice, use a disposable scalpel to gently cut across the slice next to the DG, parallel to the CA1 cell layer (Figure 1). Then cut the remaining slice vertically to remove the CA3 subfield and the subiculum. Cut a corner of the slice to help identify the top surface of the tissue.

NOTE: In case no viral delivery of protein of interest is required, the entire tissue slice can be fixed after the media is removed with a gentle rinse of ice-cold buffer. This is then followed by cutting out the CA1.

CRITICAL STEP: Keep track of the topside of the tissue in order for proper sectioning of the grids later on.

3. Carefully remove the tissue from the membrane using the backside of the scalpel. Use a Pasteur pipette to gently transfer the tissue to a 12-well plate containing 0.1 M phosphate buffer.
4. Remove the buffer in the well from the plate and add 500 μl - 1 ml of ice-cold fixative (pH 7.3) comprised of the following: 0.1% picric acid, 1% paraformaldehyde, and 2.5% glutaraldehyde in 0.1 M phosphate buffer. Incubate for 2 hr at 4 °C.

NOTE: Picric acid may be explosive when dry. Keep it wetted with water in a container tightly closed. Store in a dry and well-ventilated place.

5. Remove the fixative from the well and wash samples 3 times (20 min each) in 0.1 M phosphate buffer.
6. Incubate for 40 min in 1% tannic acid (w/v) in 0.1 M maleate buffer, pH 6.0.
7. Rinse twice (20 min each) in maleate buffer.
8. Incubate for 40 min in 1% uranyl acetate (w/v) in maleate buffer in the dark. Uranyl acetate is sensitive to light and is radioactive. Cover the beaker with Parafilm when dissolving and store any unused buffer in dark at 4 °C.
9. Rinse twice (20 min each) in maleate buffer.
10. Incubate for 20 min in 0.5% platinum chloride (w/v) in maleate buffer.
11. Rinse twice (20 min each) in maleate buffer. Store the samples at 4 °C until they are ready for further processing.

2. Dehydration and Embedding

Propylene oxide is carcinogenic. Avoid vapors by working with it in a fume hood. Use absolute, 100% pure ethanol that contains no trace of water. Dilute this absolute ethanol to produce different concentrations for dehydration.

Day 2

1. Incubate for 5 min in 50% ethanol and then for 5 min in 70% ethanol.
2. Incubate for 15 min in freshly prepared 1% p-phenylenediamine (PPD) in 70% ethanol.
3. Rinse three times in 70% ethanol.
4. Incubate for 5 min in 80% ethanol and then for 5 min in 95% ethanol.
5. Incubate in 100% ethanol twice 5 min each.
6. Prepare glass vials with screw caps and make sure they are clean and completely dry. Transfer slices to glass vials and label each vial simply and clearly.
7. Incubate for 5 min in 1:1 ethanol:propylene oxide.
8. Incubate in 100% propylene oxide twice 5 min each.
9. Add resin (Epon) to each vial to make a 1:1 mixture with propylene oxide. Gently mix for 2 hr. Do not shake the vials too rigorously to prevent bubbles that could interfere with embedding.
10. Add resin to make a 3:1 mixture with propylene oxide, and mix for 2 hr.
11. Transfer to 100% resin and incubate O/N.

Day 3

12. Sandwich samples between strips of ACLAR plastic and cure for 24 hr at 60 °C.

3. Sectioning and Mounting on Grids

Day 4

1. Cut semi-thin (0.5 μm) sections and stain with 1% toluidine blue + 1% borax to determine the correct orientation of samples.
2. Cut ultrathin sections (60 nm) and mount on nickel grids, one section per grid.

4. Immunohistochemistry

All buffers and water should be filtered before use.

Day 5

1. Place a drop (~50 μl) of 1% Tween-20/phosphate buffer (T/PB), pH 7.5, on a piece of clean Parafilm on a flat surface. Gently pick up the grid with clean forceps by the edge and float it on the buffer, section down, and incubate for 10 min at RT.
2. Drain excess liquid from the grid by placing section side up on to filter paper and then float on 50 mM glycine in T/PB for 15 min at RT.

CRITICAL STEP: To avoid excessive 'carry-over' of solutions from one solution droplet to the next during the incubations, drain excess liquid by gently touching the edge of the grid onto #1 filter paper between each solution change. Also remove any solution trapped between the arms of the EM forceps holding the grid by gently wicking with a sliver of filter paper between the forceps arms while ensuring the sections do not dry out completely.

3. Dry the grid with filter paper and then float on blocking solution containing 2.5% BSA and 2.5% serum from the animal of the secondary antibody in T/PB for 30 min at RT.
4. Incubate with primary antibody (in T/PB) at RT or O/N at 4 °C. The optimal time and temperature of incubation, as well as antibody concentration, need to be experimentally determined.
5. Wash the grid three times (2 min each) with T/PB.
6. Incubate with secondary antibody (1:20 anti-rabbit or anti-mouse coupled to 10 nm gold) for 1 hr at RT.
7. Wash three times (2 min each) with T/PB.
8. Postfix with 2% glutaraldehyde in T/PB for 5 min.
9. Wash three times (2 min each) with T/PB and then three times (2 min each) with filtered water.
10. Incubate with 2% uranyl acetate in water for 10 min.

While the grid is staining, prepare a CO₂-free chamber by placing a piece of Parafilm in the center of a glass Petri dish. Then place 4-6 pellets of NaOH in the dish around the Parafilm to absorb CO₂ in the air. Keep the top closed for a few minutes. Use a Pasteur pipette and quickly transfer a small volume of Reynold's lead citrate solution to the Parafilm in the glass Petri dish. Open the top just enough to insert the Pasteur pipette to minimize the re-introduction of CO₂ into the chamber.

NOTE: To make Reynold's lead citrate solution, boil 100 ml of deionized water in a microwave oven and let cool in an airtight container. In a 50 ml volumetric flask with stopper, mix 1.33 g lead nitrate, 1.76 g sodium citrate and 30 ml of the boiled water by shaking vigorously for 1 min. Then shake intermittently for 30 min. To this cloudy solution, add 8 ml of 1 M sodium hydroxide and slowly invert the flask a few times. Solution will become clear. Bring the solution to 50 ml with the boiled water. Store the solution tightly sealed. If precipitate appears, discard and make a new one.

11. In three smaller beakers prepare warm, freshly boiled deionized water. Wash off the uranyl acetate by dipping the grid in the first beaker and gently swirl it around for 30 sec. Repeat this step in the other two beakers.
12. Open the top of the glass Petri dish just enough to place the grid on the drop of Reynold's lead citrate solution. If possible, wear a mask while doing this to avoid breathing on lead citrate and to prevent formation of precipitate. Incubate for 10 min.
13. Open the top just enough to remove the grid. Avoid breathing on the lead citrate. Wash the grid three times by dipping it sequentially in three beakers with warm, freshly boiled distilled water. Let the grid dry on a piece of filter paper, section up. The sample is now ready for the electron microscope.

Results

Figure 2B shows an example of the distribution of endogenous Ng molecules in dendritic spines of CA1 hippocampal pyramidal neurons. Nickel grids with ultrathin (60 nm) tissues containing CA1 region of the hippocampus (as seen in Figure 2A) were covered in 1% T/PB, 50 mM glycine, then blocked with 2.5% BSA and 2.5% serum prior to incubation with anti-Ng antibody. After washing with T/PB, grids were then covered in anti-rabbit secondary antibody coupled to 10 nm gold. Finally, grids were w...

Discussion

In this protocol, we have adopted the Phend and Weinberg method for brain and spinal cord tissues to study dendritic spines in rat hippocampal slice cultures. Dendritic spines in the hippocampal CA3-CA1 area are delicate structures containing a vast variety of proteins that play important roles in regulating neuronal functions. The presented method provides a balanced approach for achieving enhanced antigenicity while maintaining good ultrastructural preservation (Figure 2A), permitting reasonable...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
60 x 15 mm polystyrene Petri dish	Falcon	351007
Disposable scalpel	EXELINT	29552
Cell culture inserts	Millipore	PICM03050
10 nm Goat-anti-rabbit gold	Electron Microscopy Sciences	25108
Anti-Neurogranin antibody	Millipore	AB5620
100% Picric acid	Electron Microscopy Sciences	19550
96% Paraformaldehyde	Acros Organics	AC41678-0030
25% Glutaraldehyde (EM grade)	Sigma	G5882
Uranyl acetate	Electron Microscopy Sciences	22400
p-Phenylenediamine	Sigma	P6001
Platinum (IV) chloride	Sigma	379840
Tannic acid	Electron Microscopy Sciences	21710