Materials and Reagents

Surgical blade (FEATHER Safety Razor, catalog number: No. 23 )
1.5 ml microcentrifuge tube
Pipette tips
Conventional needles PrecisionGlide 23 G 1 in. (Fisher Scientific, catalog number: 14-826-6B)
Manufacturer: BD, catalog number: 305193 .
BD disposable syringes (BD, catalog number: 309628 )
Cover Glass No. 1 ½ 22 x 30 mm (e.g., Corning, catalog number: 2850-22 )
Microscope slides (e.g., Fisher Scientific, catalog number: 12-552-3 )
Flies expressing sarcomere fluorescent markers (e.g., Zasp52-GFP, Table 1)
Custom-made 3.5% agar-filled 60 x 15 mm Petri dish plate (BioShop, catalog number: AGR003 )
Phalloidin-Tetramethylrhodamine B isothiocyanate (TRITC) (Sigma-Aldrich, catalog number: P1951 )
Phalloidin-Fluorescein Isothiocyanate (FITC) (Sigma-Aldrich, catalog number: P5282 )
Mounting media ProLong Gold Antifade Mountant (Thermo Fisher Scientific, Invitrogen^TM, catalog number: P36934 )
Magnesium chloride (MgCl₂)
Ethylene glycol-bis-tetraacetic acid (EGTA) (Sigma-Aldrich, catalog number: E3889 )
Adenosine triphosphate (ATP) (BioShop, catalog number: ATP007 )
1,4-Dithiothreitol (DTT) (BioShop, catalog number: DTT001 )
cOmplete^TM, EDTA-free Protease Inhibitor Cocktail (Roche Diagnostics, catalog number: 11873580001 )
Glycerol (BioShop, catalog number: GLY001 )
Triton X-100 (BioShop, catalog number: TRX777 )
8% paraformaldehyde aqueous solution glass vial (Electron Microscopy Sciences, catalog number: 157-8 )
Sodium chloride (NaCl)
Potassium chloride (KCl)
Sodium phosphate dibasic (Na₂HPO₄)
Potassium phosphate dibasic (K₂HPO₄)
Relaxing solution (see Recipes)
Relaxing-Glycerol solution (see Recipes)
8% paraformaldehyde (see Recipes)
10x phosphate buffered saline (PBS) (see Recipes)
1x PBS, 0.1% Triton X-100 (PBST) (see Recipes)

Equipment

CO₂ Flypad, standard size (8.1 x 11.6 cm) (Genesee Scientific, Flystuff, catalog number: 59-114 )
Blade handle for surgical blade (FEATHER Safety Razor, catalog number: No. 4 )
Glass Petri dish; 60 x 15 mm (VWR, catalog number: 89000-770)
Manufacturer: DWK Life Sciences, KIMBLE^®, catalog number: 2306-10010 .
Stereo microscope (Leica Microsystems, model: Leica MS5 )
Dumont #5 forceps (Fine Science Tools, catalog number: 11251-30 )
Incubator set to 25 °C
P2, P20, P100, and P1000 Micro Pipettes (e.g., Gilson, catalog numbers: F144801 , F123600 , F123615 and F123602 )
Platform mixers (e.g., Speci-Mix Test Tube Rocker) (Thermo Fisher Scientific, Thermo Scientific^TM, catalog number: M71015Q )
Medium-sized pointed brush
Standard fly husbandry equipment

Software

Fiji (https://fiji.sc/)
R Statistics package (https://www.r-project.org/)

Procedure

Prepare flies to dissect
1. Anesthetize flies using standard CO₂ pads. Use 5-10 days old flies.
2. Transfer flies to a custom-made agar plate using a fine brush. Concentrate all flies in the center of the plate.
3. Add 100 µl of Relaxing-Glycerol solution (see Recipes) to the flies using a previously cut pipette tip.
Cut half-thoraces
The main goal of this step is to split the thorax into two symmetric halves (Figure 1 and Video 1). This step is relatively easy, however take note not to crush the muscles. If the blade is dull, replace it with a new blade.
1. Use two fine forceps to remove the abdomen. Leave only the thorax and the head.
2. Place and align the thorax with the dorsal side facing your eyes.
3. Using a scalpel blade knife and a fast slicing movement cut the thorax in two symmetrical halves. Cut the head out if still attached to the thorax.
4. At this point the IFM should be visible under the dissection microscope.
5. Take the half-thoraxes with forceps and place them in a 1.5 ml microcentrifuge tube.
6. Add 100 µl of Relaxing-Glycerol buffer.
  
  Figure 1. Dissection of the thorax into two halves. A-D. Snapshots from Video 1 showing critical dissection steps. A. Place the thorax with the dorsal side up; B. Gently touch the thorax to make sure the blade is positioned exactly at the middle and that the thorax will not move. C. Aim the tip of the blade at the head/thorax attachment site (arrowhead); D. With a sharp push cut the thorax; E. Representative thorax divided in two. Muscle fibers can be observed at this stage (red asterisks). Scale bars = 200 µm.
  
  Video 1. Dissection of the thorax into two halves. Dissection process recorded directly from the stereomicroscope. Snapshots of this video are presented in Figure 1.
Remove mitochondria
The mitochondria can be completely removed from the sample by an overnight incubation in Relaxing-Glycerol buffer at -20 °C without agitation (Figure 2). Removal of mitochondria makes imaging and staining easier. If mitochondria must be preserved, skip this section.

Figure 2. Effect of glycerol extraction on mitochondria. A and B. Representative IFM expressing a mitochondrial:GFP protein and a mCherry-tagged Zasp52. A. In non-glycerinated IFM mitochondria can be detected; B. Mitochondria cannot be detected in glycerinated IFM. Scale bars = 10 µm.
Paraformaldehyde fixation
1. Add 150 µl of 8% paraformaldehyde (see Recipes) to the tube that contains the thoraces in relaxing buffer. The final paraformaldehyde concentration should be around 4%. A 4% formaldehyde fixation solution can also be used.
2. Incubate at room temperature for 45 min with slow agitation (~50 rpm).
3. Wash by replacing fixing solution with PBST (see Recipes). Incubate in PBST for 15 min under slow agitation (~50 rpm). Repeat this step three times. Thoraces quickly sink to the bottom of a microcentrifuge tube.
Dissection of muscle fibers
The main goal of this step is to separate the DLM fibers from the carcass without damaging them (Figure 3 and Video 2). Video 2 shows a detailed real-time dissection, see it before attempting to dissect the muscle fibers to get an idea of the overall process.
1. Transfer the thoraces to a glass Petri dish using a previously cut pipette tip.
2. Add enough PBST to just cover the tissue (excess liquid will make dissection challenging).
3. Connect the needles to the syringes to make two dissection needles.
4. Use two dissection needles to dissect the muscle fibers from the carcass (for details see Figure 4 and Video 2).
5. Separate the fibers as much as possible without tearing the fibers apart.
6. Transfer the fibers using a previously cut pipette tip back to a 1.5 ml microcentrifuge tube with PBST.
  
  Figure 3. General description of the IFM. A. Schematic view of a longitudinal section of a thorax, showing the position of the IFM. Two types of muscles compose the IFM, the dorsal longitudinal muscles (DLM, orange) and the dorsal ventral muscles (DVM, green). The goal of this protocol is to dissect the DLM. B. Representative isolated DLM, orange asterisks indicate individual DLM fibers (see also Video 2). Scale bar = 200 µm.
  
  Figure 4. Step-by-step guide to dissect DLM fibers. A-F. Snapshots from Video 2. A. Locate and hold the half-thorax using the left needle. Place the needle just below the DLM and above the legs. B. Make a small incision at the DLM posterior attachment site. C. Make a long cut above the DLM to completely separate the muscle fibers from the dorsal cuticle. D. Move the right needle to the ventral DLM (near the left needle). E. With the right needle push the DLM fibers out of the thorax. F. Isolated DLM can be further separated into individual fibers. G. Cartoon illustrating the dissection process. Dotted red lines mark the dissection needles. Note the small amount of liquid (~100 µl) in the dissection plate to prevent floating of half-thoraces. Scale bars = 200 µm.
  
  Video 2. Dissection of the DLM fibers. Stereomicroscope-recorded video showing the steps required for the correct dissection of the DLM fibers. Snapshots of this video are also presented in Figure 4.
Phalloidin staining and mounting
1. Replace the PBST solution with detergent-free PBS. Be careful not to aspirate the fibers.
2. Add FITC- or TRITC-phalloidin (1:1,250) in triton-free PBS.
3. Incubate at room temperature for 1-2 h with slow agitation (~50 rpm).
4. Wash three times with PBST under slow agitation (~50 rpm).
5. Transfer the muscle fibers to a glass slide.
6. Dry the samples using a Kimwipe.
7. Add a drop of mounting media, compatible with fluorescent labelling. Cover with coverslip compatible with confocal microscopy.
8. Store at 4 °C.
Confocal microscopy
1. Locate the muscle fibers under brightfield at 20x magnification.
2. Change the objective to 63x/1.4 NA and adjust the focus using brightfield to protect the sample.
3. Scan the muscle fiber using the confocal software, align the fiber and take a full resolution scanning image at 9x magnification 1,024 x 1,024 pixels.
4. Take several 9x images at different locations within the muscle fiber to get an accurate representation of the phenotype, especially when dealing with tissue from a mutant.

Data analysis

Image analysis
1. Images obtained from this protocol can be used to identify sarcomere phenotypes. Direct comparison of images from mutant muscles and control muscles should be enough to detect even subtle sarcomeric phenotypes. As healthy IFM are very regular structures (see Figure 5 for control examples), the mutant phenotypes can be scored by directly counting the number of normal sarcomeres in control and mutant muscles.
2. When counting phenotypes exclude all sarcomeres that appear out of focus, do not have well-defined actin staining, or have more than one Z-disc per sarcomere. These are indications of artifacts often caused by overlapping myofibrils. Actin staining should always be used as counterstain when counting phenotypes.
  
  Figure 5. Representative examples of IFM confocal images. A. Zasp52-mCherry stained with phalloidin to visualize actin thin filaments. B. GFP-Actinin (GFP-Actn) and Zasp52-mCherry show a clear Z-disc colocalization. C. Flightin-GFP (Fln-GFP) marks the part of the A-band where the myosin heads are present. Fln does not colocalize with Zasp52-mCherry. D. Obscurin-GFP localizes to the M-line. Scale bars = 10 µm.
Semi-automatic Z-disc or M-line measurements
The fluorescent signal from Z-disc, M-line, or A-band proteins can be used to automatically detect individual particles (Figure 6). This approach is useful when very precise measurements of a large number of sarcomeres are required (e.g., estimating Z-disc size). Video 3 shows a screen-recorded version of this protocol.
1. Open the image with the Fiji package of ImageJ (Schindelin et al., 2012).
2. Open the threshold menu, under the Image/Adjust menu. Select ‘Dark background’ and ‘Auto’.
3. Under ‘Analyze’ menu, select ‘Set measurements’. Check the Area, Standard deviation, Bounding rectangle, Shape descriptors, Mean gray value, and Display label boxes.
4. Under ‘Analyze’ menu, select ‘Analyze particles’
5. Adjust the size value (0.3-1) and/or circularity to exclude undesirable objects (e.g., protein aggregates in mutant muscles).
6. Select ‘outlines’ in the drop-down menu.
7. The measurement and outline of the image should appear. If the outline contains undesirable objects, adjust the parameters in the ‘Analyze particles’ menu.
8. Save the results.
9. Analyze the results with any statistical software (e.g., R statistical programming language).
  
  Figure 6. Automatic measurement of individual Z-discs. A. Confocal image of Zasp52-mCherry; B. Thresholded image showing in black the pixels to be included. The thresholding step uses signal intensity as an inclusion criterion. C. Result from running the ‘analyze particles’ step. This step uses size and shape as inclusion criteria. For additional details see Video 3. Scale bars = 10 µm.
  
  Video 3. Automatic detection and measurement of individual Z-discs. Screen-recorded video showing the procedure in Fiji to obtain individual Z-discs and measure some properties of the isolated discs. First, a thresholded image is made using the automatic threshold tool. Then, individual Z-disc are obtained using the ‘Analyze particles’ tool. The particle size and the circularity options may be adjusted to optimize the procedure.

Notes

Antibody stainings
Antibody stainings work very well with IFM. However, they do not seem to work perfectly when this protocol is used. If antibodies should be used, the fixation and the dissection steps must be inverted. That is, after the overnight incubation instead of fixing the tissue first dissect the IFM fibers as shown in Video 2. Then wash the samples 3 times with Relaxing solution and then proceed to the fixation step. While this greatly enhances antibody stainings, unfixed DLM muscles are harder to dissect than pre-fixed fibers.
Useful fluorescent markers
This protocol works best when fluorescent markers are available. Many different stocks expressing GFP-tagged proteins are available from common stock centers. Table 1 shows a list of useful GFP stocks for IFM imaging. To complement the existing GFP stocks we made Zasp^{52MI02908-mCherry}by integrating an mCherry cassette in frame with the Zasp52 sequence. This new Zasp^{52MI02908-mCherry} allele recapitulates Zasp52 expression and localization (Figures 2, 5, and unpublished observations).

Table 1. Useful fluorescent markers

Note: * Not fully described, available upon request.
Confocal images
1. All images must be acquired at the same resolution, magnification, and orientation. This allows direct comparison of images and saves time when arranging figures. We recommend a 20 x 20 µm square image, scanned at 1,024 x 1,024 pixel resolution and averaged 3 times.
2. Scan the muscle fiber as close to the surface as possible. Phalloidin has limited infiltration into the IFM fibers. Thus, actin staining will be sharp at the surface but fuzzy in the interior. More importantly, IFM phenotypes may slightly vary between the surface and the interior of the muscle fibers.

Recipes

Relaxing solution
1. Dissolve in 40 ml of ddH₂O:
  1,000 µl (final concentration is 20 mM) of sodium phosphate buffer stock solution 1 M pH 7.0
  500 µl (final concentration is 5 mM) of 0.5 M MgCl₂
  2.5 ml (final concentration is 5 mM) of 0.1 M EGTA
  2.5 ml (final concentration is 5 mM) of 0.1 M ATP stock solution
  250 µl (final concentration is 5 mM) of 1 M DTT
2. Add 1 tablet of protease inhibitors (cOmplete^TM, EDTA-free Protease Inhibitor Cocktail, Roche)
3. Adjust volume to 50 ml with ddH₂O
4. Store at -20 °C
Relaxing-Glycerol solution
Note: Prepare Relaxing-Glycerol solution based on York Modified Glycerol solution (Peckham, 1990).
1. Mix the relaxing solution with glycerol 1:1 (v/v)
2. Add 0.5% Triton X-100
3. Store in 10 ml aliquots at -20 °C
8% paraformaldehyde
1. Open the 8% paraformaldehyde aqueous solution glass vial (Electron Microscopy Sciences)
2. Divide the vial content into 400 µl aliquots and store them at -20 °C. Aliquots can then be stored for more than 6 months
10x phosphate buffered saline (PBS)
1. Dissolve in 800 ml ddH₂O
  80 g of NaCl
  2.0 g of KCl
  14.4 g of Na₂HPO₄
  2.4 g of K₂HPO₄
2. Adjust pH to 7.4.
3. Adjust volume to 1 L with ddH₂O
4. Dilute to 1x PBS with ddH₂O
1x PBS, 0.1% Triton X-100 (PBST)
Add 500 µl of Triton X-100 to 500 ml of 1x PBS

Acknowledgments

This protocol is adapted from the work of others. The original IFM dissection technique was described and distributed by Belinda Bullard and John Sparrow. We thank Anja Katzemich for technical advice, and the CIAN imaging facility for help with confocal microscopy. This work was supported by operating grant MOP-142475 from the Canadian Institutes of Health Research. We have no conflict of interest to declare.

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