Immunofluorescent labelling of adult mouse neuromuscular junctions

Dissection and freezing

Reagents and equipment required:

1. Rapidly dissect the extensor digitorum longus (or other) muscle from tendon to tendon.
2. Remove excess connective tissue and blot the muscle on paper towel if necessary.
3. Place the muscle in a 24 well plate and fix in ice cold 4% PFA for 10 minutes.
   1. Ensure the muscle has not contracted or coiled up, stretch it out across the base of the well.
4. Wash 3 x 10 min in PBS. 
   1. Complete all washes at room temp on a platform rocker.
5. Incubate the muscle in 30% sucrose (made in PBS) for approx. 24h at 4C°.

From this point, you can opt to dissociate the entire muscle, flattening it out for a whole mount approach or freeze the sample for cryosectioning and NMJ labelling of 100 µm thick serial sections.

Wholemount approach

1. Place the muscle in a petri dish containing PBS, remove connective tissue and manually dissociate myofibres using a pair of fine point (#5) forceps by gently teasing apart the muscle fibre bundles. The muscle will begin to spread and flatten out as more fibres are teased apart.
   1. Begin teasing fibres apart from either the proximal or distal end of the muscle, at the tendon.
2. Wash the muscle 2 x 10 min in PBS to remove sucrose residue.
3. Transfer the muscle to a 24 or 48 well plate to reduce volumes. Volumes will depend on well size, ensure tissue is submerged (300 ul for 48 wells).

Cryosectioning approach

Before you start

• Pour approx. 40 ml of isopentane into the float boat and gently place the boat onto LN_2.
  1. Vapour will come off the LN₂ rapidly.
  2. Always leave the lid off the dewar when the float boat is inside.
  3. Don’t allow the isopentane and LN₂ to mix.
  4. The isopentane will eventually freeze into a solid block. 

1. Wash the muscle 2 x 10 min in PBS to remove sucrose residue
2. Squeeze OCT freezing compound into a cryomold up to the brim. Remove any bubbles.
3. Blot off excess PBS
4. Embed the muscle longitudinally, directly into the OCT, in the cryomold ensuring the muscle is pressed down flat against the bottom of the cryomold.
5. Gently place the cryomold on top of the solid frozen isopentane inside the LN₂ dewar.
   1. Wait until the OCT has frozen solid usually 30-60 seconds.
6. Place the cryomold on dry ice until storing in the -80°C freezer long term.

Cryosectioning

Reagents and equipment required:

Specific temperatures will vary based on each individual cryostat, optimise as necessary.

1. Set the chamber temperature to -20°C and the knife temperature 1-2 degrees higher than that (ie -19 or -18°C).
2. Transport the cryomolds to the cryostat on dry ice.
3. Place the cryomolds into the chamber and equilibrate for 20-30 minutes.
4. Pop the sample out of the cryomold, be sure to maintain the orientation of the block.
5. Mount the block onto the chuck using OCT, allow to set.
6. Section according to facility SOP.
   1. Trim the block sufficiently to ensure you are deep enough into the muscle. Trimming at 20 µm is advised. 
   2. Ensure the pink colour of the muscle is clear and no white/opaque OCT is present on the cutting face of the sample. This will ensure you are deep enough into the sample.
7. Serial section the muscle, longitudinally at 100 µm.
8. Using chilled forceps, gently place the section onto a slide.
   1. While sectioning, keep slides outside of the refrigerated chamber.
   2. When it comes time to place the section on the slide, hold the slide inside the chamber and then place the section directly onto the slide.
   3. The temperature difference between the slide and the section will allow for easy adherence between the section and the slide. 
   4. From one EDL muscle, you should be able to recover a minimum of 6 sections and place 3 serial sections per slide.
9. Allow slides to dry at room temperature before storing at -20°C.
   1. Storage at -80°C is also suitable.

Immunofluorescent label

Reagents and equipment required:

* Alternate wavelengths are also suitable, user at your own discretion

The IF can be performed on either whole muscles (teased apart and flattened) or 100 µm thick sections. If using whole muscles, perform all steps in a well plate.

Day 1

1. If using sections, on the day, leave slides at room temp for 2 hours before beginning the protocol.
2. Perform all washes on the rocker.
3. 3 x 5 min washes in PBS.
4. Permeabilise by incubating in 2% Triton X in PBS (30 min at RT).
5. If using slides, place slides in a humidity chamber.
6. Block in 10% goat serum and 1% Triton X in PBS (30 min at RT).
   1. If serum is not available use 4% BSA.
7. Incubate in primary antibody solution of SV2 (1:100) and 2H3 (1:50) in blocking solution overnight at 4°C.
   1. For whole muscles, longer incubation periods in primary antibody may be necessary, this will depend on the size of the tissue. Incubation over a period of multiple nights is suitable. Optimise as per your conditions.

Day 2

1. 3 x 15 min washes in PBS. 
2. Incubate in secondary antibody solution of AF594 goat – anti mouse (1:250) and 488 conjugated α-Bungarotoxin (1:500) in PBS for 2 hours at room temp in the dark.
3. 3 x 5 min washes. 
4. Cover slip with round 13 mm No 1.5 coverslips and leave to dry at 4°C overnight in the dark.
   1. Use coverslips that are large enough to cover the entire specimen, optimise accordingly.
   2. Note – if mounting an entire (flattened muscle) you may need larger coverslips to ensure the entire muscle is effectively mounted. Because the specimen is thicker than a usual section, you may need to pipette more Mowiol under the cover slip to ensure there are no air bubbles.
5. Seal the coverslip with nail varnish if necessary.

Confocal microscopy

Use a confocal microscope to image NMJs. The imaging conditions will vary across different microscopes however, the following information is universal:

1. Capture a low magnification image of the entire muscle (if possible) and use this as a map to capture all regions of interest across the muscle.
2. Capture as many NMJs as possible ensuring to Z stack through the entire sample.
   1. Ensure z-steps are not too far apart, less than 2.5 µm is suggested.
   2. If imaging sections (100 µm), ensure you image at least 3 serial sections.
3. 63x immersion capture is required for aNMJ morph analysis https://doi.org/10.1098/rsos.200128.
4. Simple manual scoring can be performed on 20x images using ImageJ. 
5. Maximally project Z stacks to score images.