Immuno-correlative light and electron microscopy (iCLEM) using TEM

Dissect out the tissue of interest (example, forebrain of Tg(proliferating cell nuclear antigen:GFP) transgenic adult zebrafish with olfactory bulbs attached for tissue orientation) on a teflon plate or dental wax sheet using fine forceps and a scalpel blade, at room temperature and place in fixative, either 2% paraformaldehyde, 0.2% glutaraldehyde in 0.1M phosphate buffer (PB) (pH 7.4) or 4% paraformaldehyde in 0.1M PB (pH 7.4). Keep the tissue submerged in the fixative at all times.

Always perform fixation using a fume hood and wear appropriate personal protective equipment (PPE).

Place the tissue in 5 ml tubes containing the fixative used in Step 1 (2% paraformaldehyde, 0.2% glutaraldehyde in 0.1M PB (pH 7.4) or 4% paraformaldehyde in 0.1M PB (pH 7.4)) and fix overnight at 4 degrees Celsius on a rotor. For brain samples, add 4% sucrose to the fixation solution.

Samples can be stored in 1% paraformaldehyde in 0.1M PB at 4 degrees Celsius until further processing.

Remove the fixative and wash with phosphate buffered saline (PBS), 3 x 10 mins.

Wash with 0.15% glycine in PBS for 10 mins.

Infuse tissue with pre-warmed 6% gelatin in 0.1M PB for 1 hour at 37 degrees Celsius, agitating. Use a heat block to maintain the temperature.

Infuse tissue again with pre-warmed 6% gelatin in 0.1M PB for 1 hour at 37 degrees Celsius, agitating.

Place gelatin infused tissue in plastic embedding moulds containing pre-warmed 6% gelatin in 0.1M PB at 37 degrees Celsius, and solidify gelatin blocks by cooling at 4 degrees Celsius for 60 mins.

Alternatively, the flat embedding method can be used to minimise gelatin block shrinkage following sucrose infiltration. Add 6 percent pre-warmed gelatin in 0.1M PB to a petri dish and solidified at 4 degrees Celsius for 1 hour. Place the tissue on top in a layer of 6 percent gelatin pre-warmed to 37 degrees Celsius. Solidify gelatin at 4 degrees Celsius for 1 hour.

Place the gelatin infused tissue blocks on a teflon plate or dental wax sheet and cut off excess gelatin from around the tissue using a razor blade or scalpel.

For flat embedded samples, cut around the tissue using a razor blade or scalpel and remove the gelatin embedded sample from the petri dish using a small spatula. Cut off excess gelatin using a razor blade or scalpel.

Fix the gelatin infused tissue blocks in 0.2% paraformaldehyde in 0.1M PB for 30 mins at 4 degrees Celsius, rotating.

Wash tissue blocks with 0.1M PB, 3 x 10 mins, agitating.

Infuse tissue blocks with 2.3M sucrose in 0.1M PB for 2 days at 4 degrees Celsius, rotating.

Mount each block on a clean aluminium bullseye pin. Position tissue in the correct orientation and remove the excess sucrose with strips of filter paper.

Freeze each sample block by gently submerging the pins in liquid nitrogen. For large blocks, cool the sample pin in the chamber of a cryo-ultramicrotome (-100 degrees Celsius) for 10 mins prior to freezing in liquid nitrogen. Pins should be mounted and frozen one at a time.

Trim the front face and edges of the frozen tissue block cutting 50-100 μm deep on all four sides. Cut at a speed of 100 mm/sec with a 100 nm feed, at -90 to -100 degrees Celsius using a Leica UC7/FC7 cryo-ultramicrotome and Diatome cryotrim 20 knife.

To check the region of interest:

Cut semi thin sections with a feed of 100-300 nm, at 3 mm/sec and at -90 to -100 degrees Celsius.

Pick up sections in a 1:1 mixture of 2% methylcellulose: 2.3M sucrose in 0.1M PB using a 3 mm stainless steel loop.

Place sections on a slide and stain with methylene blue/Azure II or toluidine blue solution for 10-20 seconds before rinsing with milliQ water and drying on a mini hotplate.

Observe sections with a wide-field light microscope. If the region of interest has not been obtained, continue trimming and repeat Step 16.

Once the region of interest has been obtained, cut 62-70 nm ultrathin sections at 0.8 mm/sec and -90 to -100 degrees Celsius using a Leica UC7/FC7 cryo-ultramicrotome and Diatome cryo immuno knife.

Pick up ultrathin sections in a 1:1 mixture of 2% methylcellulose : 2.3M sucrose in 0.1M PB using a 3 mm stainless steel loop, and place sections on 50-150 mesh copper or palladium coated copper grids with carbon-coated formvar film.

Store grids in an enclosed petri dish at 4 degrees Celsius until use.

Consecutive (serial) 62-70 nm tissue sections of the same cell type can be used for the following workflows:

1. Immunofluorescence (IF) labelling of ultrathin sections for optical microscopy using different cellular markers (Steps 20-37) and then preparation of these grids for subsequent TEM imaging to obtain ultrastructural detail (Steps 38-48).

2. Single or double-immunogold labelling of ultrathin sections and TEM imaging to validate the specificity of IF markers at high resolution (Steps 49-77).

When used in combination, these methods allow for the correlation of two fluorescent markers with two different sized protein-A-gold markers within the same cell, as described below.

Fill the wells of a 24-well plate with PBS and float grids section side down.

Use a fine tip pasteur pipette to remove enough PBS from each well so that the lid can be secured on the well-plate without disturbing the floating grids. Seal the lid with parafilm.

Put the well-plate containing the grids in an oven set to 50-60 degrees Celsius for 1 hour to remove the 2% methylcellulose : 2.3M sucrose and 6% gelatin from the tissue sections.

Place a sheet of parafilm on a clean flat bench or a glass board using a small amount of water underneath to keep the film flat.

For the following steps, use a perfect loop or fine forceps to transfer grids from one drop of solution to another, section side down. Use approximately 100-200 μl drops for rinsing solutions and 5 μl drops for antibodies on a clear parafilm surface. Centrifuge all antibodies for 30 seconds using a benchtop microcentrifuge prior to use.

Quench aldehydes with 0.15% glycine in PBS, 5 x 2 mins.

Block with 1% bovine serum albumin (BSA) in PBS for 5 mins.

Incubate with both primary antibodies diluted in 1% BSA/PBS for 45 to 60 mins in a dark moist chamber, at room temperature.

Example: Mouse anti-glutamine synthetase (1:500 dilution), to label glial cells.

Example: Biotinylated anti-eGFP (1:300 dilution), to label green fluorescent protein labelled proliferating cell nuclear antigen (PCNA).

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Optionally incubate with a bridging antibody diluted in 1% BSA/PBS for 30 mins at room temperature.

Example: Rabbit anti-biotin (1:10,000 dilution)

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Incubate with both secondary antibodies diluted in 1% BSA/PBS for 45 mins in a dark moist chamber, at room temperature.

Example: Goat anti-mouse AlexaFluor-555 (1:300 dilution).

Example: Goat anti- rabbit AlexaFluor-488 (1:300 dilution)

Rinse with PBS, 5 x 2 mins.

Rinse with distilled water, 4 x 2 mins.

Incubate with Hoechst nuclear stain (1 μM in distilled water) for 20 mins.

Rinse with water, 4 x 2 mins.

Place grid section side up in a drop of 50% glycerol in water. Ensure the grid is completely covered in glycerol with no air bubbles present.

Place grid section side down on a 30 mm glass bottom culture dish and cover grid with a glass coverslip.

Acquire a fluorescent montaged z-stack of 62-70 nm ultrathin sections using an inverted widefield fluorescent microscope.

Example: Leica AF6000LX with a DFC 350FX camera, and a 40x 0.6 NA dry or 63x 1.3 NA glycerol objective.

After imaging, prepare the grids for transmission electron microscopy as follows:

Add distilled water to the 30 mm glass bottom culture dish and agitate gently until the coverslip floats.

Remove the coverslip and lift the grid out using fine forceps.

Carefully remove water from the backside of the grid by placing filter paper at the grid edge.

Put a drop of distilled water on a sheet of parafilm and place the grid on the drop. The grid should float.

Rinse with distilled water, 6 x 2 mins. Water drops should be placed on a clean parafilm sheet.

Stain the grid in a drop of 2% Uranyloxalate (pH 7.0) for 5 mins at room temperature.

Rinse once with distilled water.

Cover a glass petri dish with parafilm using a small amount of water under the film to keep it flat. Place the dish on ice and add 3 drops of filtered 2% methylcellulose : 4% uranyl acetate (9 ml : 1 ml) (pH 4.0) to the surface.

Float the grid briefly in the first two drops of methylcellulose/uranyl acetate, then transfer to the third drop and leave for 10 minutes.

Remove and dry the grid following the looping out method:

Clean a remanium wire loop with water and dry.

Push the loop into the methylcellulose/uranyl acetate drop and under the grid.

Lift the grid out from the drop using the loop.

Tilt the loop and grid at a 45-60 degree angle and bring the loop to a piece of filter paper.

As soon as the loop contacts the filter paper, drag it slowly along the filter paper to remove excess liquid. A thin film of methylcellulose/uranyl acetate should remain on the surface of the grid. The slower the drag, the thinner the film.

Dry the grid in the loop for 30 minutes at room temperature.

Remove the grid from the loop using fine forceps, taking care not to tear the methylcellulose/uranyl acetate film away from the grid. Store the grid in a grid box.

Place the grid in a TEM grid holder for high resolution imaging using a transmission electron microscope.

Example: JEOL 1400-Plus 120 keV TEM equipped with a high sensitivity bottom mount CMOS 'Flash' camera.

Fill the wells of a 24-well plate with PBS and float grids section side down in the solution. Use a fine tip pasteur pipette to remove enough PBS from each well so that the lid can be secured on the well-plate without disturbing the floating grids. Secure the plate lid with parafilm.

Put the plate with the floating grids in an oven set to 50-60 degrees Celsius for 60 minutes to remove the 2% methylcellulose/2.3M sucrose and 6% gelatin from the tissue sections.

Place a sheet of parafilm on the bench using a small amount of water underneath to keep the film flat.

For the following steps, use a perfect loop or fine forceps to transfer grids from one drop of solution to another, section side down. Use approximately 100-200 μl drops for rinsing and 5 μl drops for antibodies on a clean parafilm surface. Spin all antibodies and protein A-gold (PAG) for 30 seconds using a benchtop microcentrifuge prior to use.

Quench aldehydes with 0.15% glycine in PBS, 5 x 2 mins.

Block with 1% bovine serum albumin (BSA) in PBS for 5 mins.

Incubate with a primary antibody diluted in 1% BSA/PBS for 45 to 60 minutes in a dark moist chamber at room temperature.

Example: Mouse anti-glutamine synthetase (1:500 dilution), to label radial glial cells.

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Optionally incubate with a bridging antibody diluted in 1% BSA/PBS for 20 mins, in a dark moist chamber, at room temperature.

Example: Rabbit anti-mouse Ig bridging antibody (1:1000 dilution).

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Incubate with 10 nm PAG (dilution as recommended by manufacturer for specific batch) diluted in 1% BSA/PBS for 30 mins, in a dark moist chamber, at room temperature.

Rinse with 0.1% BSA in PBS, 2 x 2 mins.

Rinse with PBS, 5 x 2 mins.

Fix with 1% glutaraldehyde in PB for 5 mins at room temperature to prevent non-specific binding of subsequent PAG particles and to stabilise the immunogold labelling reaction. This step must be completed in the fume hood using appropriate PPE.

Rinse with PBS, 2 x 5 mins.

Incubate with a primary antibody diluted in 1% BSA/PBS for 45 to 60 mins in a dark moist chamber, at room temperature.

Example: Biotinylated enhanced green fluorescent protein (eGFP) (1:500 dilution).

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Optionally incubate with a bridging antibody (1:10,000 dilution) diluted in 1% BSA/PBS for 30 mins in a dark moist chamber at room temperature.

Example: Rabbit anti-biotin bridging antibody (1:10,000 dilution)

Rinse with 0.1% BSA in PBS, 5 x 2 mins.

Incubate with 20 nm PAG (dilution as recommended by manufacturer for specific batch) diluted in 1% BSA/PBS for 30 mins in a dark moist chamber, at room temperature.

Rinse with 0.1% BSA in PBS, 2 x 2 mins.

Rinse with PBS, 5 x 2 mins.

Fix in 1% glutaraldehyde in PB for 5 mins at room temperature to stabilise the immunogold-labelling reaction. This step must be completed in the fume hood using appropriate PPE.

Rinse with PBS, 2 x 5 mins.

Rinse with distilled water, 6 x 1 min.

Stain the grid in a drop of 2% Uranyloxalate (pH 7.0) for 5 mins at room temperature.

Cover a glass petri dish with parafilm using a small amount of water under the film to keep it flat. Place the dish on ice and add 3 drops of 2% methylcellulose : 4% uranyl acetate (9 ml : 1 ml) (pH 4.0) to the surface.

Float the grid briefly in the first two drops of methylcellulose/uranyl acetate (pH 4.0), then transfer to the third drop and leave for 10 mins.

Remove and dry the grid by the looping out method, described above (Step 47).

Acquire high resolution micrographs using a transmission electron microscope.

Example: JEOL-1400 Plus 120 keV TEM equipped with a high sensitivity bottom mount CMOS 'Flash' camera.