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All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Preparation
1. Tease each muscle in small fiber bundles of about 1 mm wide using two fine serrated forceps.
NOTE: It is crucial to manipulate muscles very gently with the forceps, without excessive force, to prevent tissue damage during teasing.
1. Dissociate the tibialis anterior (TA) muscle into 3 or 4 bundles depending on its size.
2. For gastrocnemius (GA), separate the medial and lateral parts of the muscle and then dissociate each part into 4-5 bundles depending on their size.
2. Immunostaining
1. Proceed with muscle fiber permeabilization: Transfer muscle bundles into 24-well plates containing 1% (v/ v) Triton X-100 in phosphate-buffered saline (PBS) and keep them under gentle agitation (50 rpm) for 1 h at RT or 5 h at 4 °C.
NOTE: Split the muscle bundles between two plates to proceed with separate immunostainings and to minimize the risk of antibody confusion. Do not split them into more than two wells (1 well/plate); otherwise, the number (N) of neuromuscular junctions (NMJs) that are representative of their general status in the analyzed muscle may be insufficient.
2. Wash the samples 3x for 5 min with PBS at RT and incubate them with a blocking solution composed of 4% bovine serum albumin (BSA) in PBS/Triton X-100 1% for 4 h at 4 °C, under gentle agitation (50 rpm).
NOTE: Do not use an aspiration pump during the washing steps but rather aspirate the solution manually with a 200 μL pipette and small-sized tips (the reference is indicated in the Table of Materials).
3. Incubate the samples overnight (O/N) at 4 °C under gentle agitation (50 rpm) with the blocking solution indicated in step 2.2 containing primary monoclonal antibodies against either neurofilament M (NF-M, 2H3, dilution 1/200) or synaptic vesicle glycoprotein 2 (SV2, dilution 1/200) to label presynaptic axon terminals or active zones, respectively.
4. Next day, wash the muscle bundles 3x for 5 min in PBS under agitation (50 rpm).
1. For confocal imaging: Incubate the muscle bundles with secondary anti-mouse antibodies conjugated with a red-emitting fluorophore (F594) (dilution 1/500) and α-bungarotoxin conjugated with a green-emitting fluorophore (α-BTX-F488) (dilution 1/1000) in PBS for 2 h at RT under agitation (50 rpm).
2. For stimulated emission depletion (STED) imaging: Incubate the muscle bundles with secondary anti-mouse antibodies conjugated with a green-emitting fluorophore (F488) (dilution 1/500) and α-bungarotoxin conjugated with a far-red-emitting fluorophore characterized by high photostability (α-BTX-F633) (dilution 1/1000) in PBS for 2 h at RT under agitation (50 rpm).
NOTE: Do not expose the samples to light during incubation to avoid photobleaching.
5. Wash the labeled muscle bundles 3x for 5 min with PBS under agitation (50 rpm) and place them on a slide with a mounting medium.
NOTE: Place a maximum of 4 to 5 muscle bundles per slide to allow sealing.
6. Add a grade #1.5 (or #1.5H) glass coverslip (0.17 mm thickness) on the top, and place cylindrical magnets on both sides of the slide to apply pressure and flatten the muscles.
7. Keep the slides protected from light O/N at 4°C. Seal the slides permanently with nail polish.
3. Image acquisition
1. Acquisitions by a confocal microscope
NOTE: Images were collected with an inverted laser scanning confocal microscope using a 63x magnitude oil immersion objective (HCX Plan Apo CS, 1.4 numerical aperture (NA)).
1. For blinded analysis, let a person not involved in the analysis code each slide with a given number. Remain blinded to the experimental groups until the quantification of NMJ parameters is complete for all samples.
2. Launch the microscope software in Configuration Mode > machine.xlhw.
3. Place the slide on the microscope stage and find the observation plane within the sample by looking under DAPI wide-field fluorescence illumination with the DAPI filter set.
4.Click on Open Project > New Projects and create a folder to store image acquisitions.
NOTE: Create a new project for each NMJ to limit folder size and prevent computer memory issues.
5. To manage acquisition parameters, click on the Acquisition tab window and set the confocal pinhole to 1.0 Airy unit and laser power to optimize the gain and offset levels for the green/F488 (α-BTX) fluorescence using a 488 nm laser at the endplate that has to be imaged (Live mode ON).
6. Next, optimize the red/F594 (NF-M or SV2) fluorescence acquisition using a laser adapted to F594 observation. In this study, a 552 nm laser was used (Live mode ON). Set the spectrum of dye emission with the following ranges for each laser: laser 405 (DAPI) from 414 to 483 nm, laser 488 (F488-α-BTX) from 506-531 nm, and laser 552 (NFM/ SV2) from 622-650 nm.
7. Collect image stacks of neuromuscular junctions in each experimental group with the same settings: image size 1024 x 1024 pixels (73.7 x 73.7 μm) at 400 Hz sampling rate, Bidirectional X ON, Zoom factor 2.5, Z-step size 0.5 μm in Z-Wide mode.
NOTE: For each NMJ, the number of slices is set to acquire the whole junction. The acquisition settings described above fulfill the Nyquist-Shanon sampling theorem. However, the user can click on the Optimize Format button, present on all recent confocal operating software, to ensure pixel size and Z-step meet the ideal Nyquist sampling rate. This action will avoid over or under-sampled images, which will cause a loss of accuracy in volume measurements.
8. Save the original file (.lif) or Z-stack images (.tif) in a folder with a name that includes the code name of the slide, the staining type, and the endplate number.
NOTE: Collect sequentially (not simultaneously) the scans using the 488 nm and 552 nm lasers (F488 and F594) to avoid crosstalk of the F488 fluorescence into the F594 channel and vice versa (bleed-through). he beam path can be configured with the Dye Assistant in the microscope software.
9. Change to the next coded slide and repeat steps 3.1.3-3.1.8 for each NMJ.
10. At the end of the session, click on Open in 3D Viewer and choose an NMJ representative of an experimental group to visualize the 3D labeling.
NOTE: This view mode will help to verify that the acquisition parameters were correct.
11. Close the microscope software, clean the objectives with lens tissues, and then turn off the system.
2. Acquisitions by STED microscopy
NOTE: Images were collected with an inverted laser scanning confocal microscope equipped with Gated STED at 775 nm using a 100x oil immersion objective (HC PL APO CS2 1.4 NA).
1. For blinded analysis, let a person not involved in the analysis code each slide with a given number. Remain blinded to the experimental groups until the quantification of NMJ parameters is complete for all samples.
2. Launch the microscope software in Configuration Mode > machine.xlhw and STED ON.
3. Click on Open project > New Projects to create a folder to store image acquisitions.
NOTE: Generate a new folder for each slide to limit folder size and prevent computer memory issues.
4. Place the slide on the microscope stage and view it under wide-field fluorescence illumination using the 488 nm laser to find the observation plane within the sample.
5. Search for a NMJ labeled with neurofilament M (NFM) or SV2 staining using the 488 nm laser with a spectral detection from 506-531 nm.
6. When a NMJ has been identified, click on Activate STED and start acquiring images in a region that contains several junctional folds using the 635 nm laser with a spectral detection from 640-750 nm.
NOTE: Be careful to the saturation look-up table during image acquisition and click the Quick LUT button to avoid overexposure (grey values >255; for 8 bit).
7. Collect the images of each experimental group with the same settings: image size 2048 x 2048 pixels (38.75 x 38.75 μm) at a 400 Hz sampling rate.
NOTE: The depletion laser (STED) power is set to 65%.
8. Save the images with a file name that includes the code of the slide.
NOTE: It is possible to click on Optimized XY Format: Set Format to obtain the best acquisition setting with STED imaging.
9. Change to the next coded slide and repeat steps 3.2.3-3.2.8. Repeat this procedure for all slides.
10. At the end of the STED microscopy session, transfer the image files to another computer and save the original files (.lif) in an external drive or server.
11. Turn off the microscope software, clean the objectives with lens tissues, and then turn off the system.