Fiber Type Identification of Human Skeletal Muscle

Summary

This protocol demonstrates single-fiber isolation from freeze-dried human skeletal muscle and fiber-type classification according to Myosin heavy chain (MHC) isoform using the dot blotting technique. Identified MHC I and II fiber samples can then be further analyzed for fiber type-specific differences in protein expression using western blotting.

Abstract

The technique described here can be used to identify specific myosin heavy chain (MHC) isoforms in segments of individual muscle fibers using dot blotting, hereafter referred to as Myosin heavy chain detection by Dot Blotting for IDentification of muscle fiber type (MyDoBID). This protocol describes the process of freeze-drying human skeletal muscle and isolating segments of single muscle fibers. Using MyDoBID, type I and II fibers are classified with MHCI- and IIa-specific antibodies, respectively. Classified fibers are then combined into fiber type-specific samples for each biopsy.

The total protein in each sample is determined by Sodium Dodecyl-Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) and UV-activated gel technology. The fiber type of samples is validated using western blotting. The importance of performing protein loading normalization to enhance target protein detection across multiple western blots is also described. The benefits of consolidating classified fibers into fiber type-specific samples compared to single-fiber western blots, include sample versatility, increased sample throughput, shorter time investment, and cost-saving measures, all while retaining valuable fiber type-specific information that is frequently overlooked using homogenized muscle samples. The purpose of the protocol is to achieve accurate and efficient identification of type I and type II fibers isolated from freeze-dried human skeletal muscle samples.

These individual fibers are subsequently combined to create type I and type II fiber type-specific samples. Furthermore, the protocol is extended to include the identification of type IIx fibers, using Actin as a marker for fibers that were negative for MHCI and MHCIIa, which are confirmed as IIx fibers by western blotting. Each fiber type-specific sample is then used to quantify the expression of various target proteins using western blotting techniques.

Introduction

Skeletal muscle is a heterogeneous tissue, with distinct cellular metabolic and contractile properties that are dependent on whether the cell (fiber) is slow twitch (type I) or fast twitch (type II). The fiber type can be identified by examining the myosin heavy chain (MHC) isoforms, which differ from each other in several ways, including contraction time, speed of shortening, and fatigue resistance¹. Major MHC isoforms include type I, type IIa, type IIb, and type IIx and their metabolic profiles are either oxidative (type I and IIa) or glycolytic (IIx, IIb)¹. The proportion of these fiber types varies in muscle type and between species. Type IIb is widely found in rodent muscle. Human muscles do not contain any type IIb fibers and consist predominantly of MHC isoforms type I and IIa fibers, with a small proportion of IIx fibers². Protein expression profiles vary between different fiber types and can be altered with aging³, exercise^4,5, and disease⁶.

Measuring cellular responses in different skeletal muscle fiber types is often overlooked or not possible due to the examination of muscle homogenates (a mix of all fiber types). Single-fiber western blot allows for the investigation of multiple proteins in individual muscle fibers⁷. This methodology has previously been utilized to produce novel and informative single-fiber characteristics that were not possible to obtain using homogenate preparations. However, there are some limitations of the original single-fiber western blot methodology, including the time-consuming nature, the inability to generate sample replicates, and the use of expensive, sensitive enhanced chemiluminescence (ECL) reagents. If fresh tissue is being used, this method is further limited due to the time constraints of needing to isolate individual fibers within a limited timeframe (i.e., 1-2 h). Fortunately, this restraint is mitigated by isolating single-fiber segments from freeze-dried tissue⁸. However, fiber collection from freeze-dried samples is limited by the size and quality of the biopsied tissue.

Fiber type identification using the dot blotting method⁹ has been significantly elaborated upon and expanded in this comprehensive protocol. Previously, it has been demonstrated that as little as ~2-10 mg of wet weight muscle tissue is adequate for freeze-drying and single-fiber MHC isoform protein analysis⁹. Christensen et al.⁹ used 30% of a ~1 mm fiber segment to detect the MHC isoform present by dot blotting, which was confirmed by western blotting. This work showed that by substituting western blotting with dot blotting, the overall costs were reduced by ~40-fold (for 50 fiber segments). Fibers were then "pooled" into type I and type II samples, which allowed for experimental replication⁹. Nevertheless, a limitation was that only two fiber type-specific samples were obtained: type I (MHCI positive) and type II (MHCII positive fibers), with type II samples containing a mixture of MHCIIa and MHCIIx^6,10. Notably, the current protocol demonstrates how pure type IIx fibers can be identified and provides a highly detailed workflow (summarized in Figure 1), including troubleshooting strategies for common protocol issues.

Protocol

Human muscle samples were obtained from the vastus lateralis from n = 3 (2 males, 1 female), aged 70-74 years old under sterile conditions using local anesthesia (Xylocaine) and a Bergstrom needle modified for manual suction^11,12. Samples were a subset of a previous study approved by the Victoria University Human Research Ethics Committee (HRETH11/221) and conducted in accordance with the Declaration of Helsinki¹³. Participants provided written, informed consent to participate in this study. Full details of all materials required for this protocol are shown in the Table of Materials. In addition, a list of troubleshooting strategies addressing common protocol issues is provided in Table 1.

1. Freeze drying

1. While keeping the biopsied muscle samples frozen on dry ice, weigh and tare an empty frozen tube on the scales.
2. Quickly weigh a minimum of 10 mg of wet weight frozen tissue. Record the weight to one decimal place.
3. Deposit the tissue into the precooled microcentrifuge tube that has had 3-4 holes pierced into the lid and place it into a small beaker with 2-3 pellets of dry ice.
4. Follow the freeze-dryer manufacturer's operating instructions.
5. Ensure that all the valves on the freeze dryer are closed and switch on the freeze dryer.
6. Use the control panel to check that the vacuum set point is programmed for optimal freeze-drying conditions for human tissue, 0.12 millibar (mBar) (Figure 2).
7. Press manual on the freeze dryer and wait until the chamber has cooled to −40 ^oC.
8. Once the chamber has reached that temperature, remove the lid, then the glass chamber, and place the beaker (with muscle sample) onto the metal stage.
9. Replace the glass chamber and lid. Close the vacuum release valve on the lid and wait until the vacuum scale light turns green to ensure the dryer is vacuum sealed.
10. Freeze-dry the muscle samples for 48 h. Once freeze drying is complete, turn off the vacuum pump by pressing the vacuum button and open the vacuum release valve on the lid.
11. Remove the lid of the chamber and collect the freeze-dried sample. Weigh the tissue and record the weight to one decimal place.
12. Calculate the percentage loss of weight; ~75% decrease in weight indicates the tissue is successfully freeze-dried (in this work, mean ± SD, 76 ± 9%, n = 11 muscle samples).
13. Press AUTO to switch off the vacuum pump and freezer. Allow the unit to reach ambient temperature.
14. Clean the cooling coils according to the manufacturer's instructions and drain accumulated liquid from the collector.
    ​NOTE: Fiber collection can commence immediately. When tissue is not being used for fiber isolation, keep the freeze-dried sample at -80 °C in a sealed container with desiccator beads.  CAUTION: Dry ice is hazardous (Class 9); refer to the MSDS for recommended safe handling.

2. Fiber collection

1. Fiber collection preparation
   1. Label a minimum of 50 microfuge tubes (Fiber 1 to 50) and pipette 10 µL of denaturing buffer to each tube.
   2. Prepare the collection area with two pairs of fine tissue dissecting forceps, lint-free tissue, a benchtop lamp, a stereomicroscope (with the black stage up), a vortex, and a benchtop centrifuge.
   3. Place the freeze-dried muscle sample onto a Petri dish lid. Place the lid on the stage of the dissecting microscope.
   4. Watch the video of single-fiber dissection. Practice the complete protocol from fiber collection to single-fiber type identification at least twice before commencing a study.
   5. Prior to fiber collection, calculate the total volume required for a fiber type sample from each biopsy as follows. For example, if the starting volume of 1 fiber = 10 µL and the volume remaining after MyDoBID is (1 fiber × 10 µL) - 1 µL used for dot blotting = 9 µL (sample volume), calculate the total sample volume required by multiplying the sample volume (µL) by the total number of western blots that will be run and double this amount to account for redundancy: (9 µL × 4 western blots) × 2 = 72 µL. Calculate the number of fibers required per typed sample by dividing the final sample volume required (µL) by the sample volume per fiber type-specific sample (e.g., 72 µL ÷ 9 µL = 8 fibers per fiber type-specific sample). For this to be achieved, collect a total of 50 fibers.
      NOTE: The sample volume required is the minimum protein concentration required to run both MyDoBID and western blotting if the amount of protein loaded is equivalent to a ~3 mm fiber (~12 µg of wet weight) for each sample (see Supplementary File 1 for details). However, this volume does not factor in whether repeat gels are required for a given protein.  CAUTION: Forceps are sharp; handle them with care to avoid the risk of skin puncture.
2. Single-fiber isolation
   1. On a small piece of paper 5 cm x 1 cm, use a ruler to draw a 1 cm line. Mark every 1 mm on that line.
   2. Insert this under the Petri dish lid as a guide to estimate the length of the fibers collected.
   3. Place the freeze-dried muscle sample under the stereomicroscope at low magnification (x 7.5). Use one pair of fine tissue dissecting forceps to hold the freeze-dried muscle in place and with the other start to separate small bundles of fibers (as seen in the video).
   4. Isolate one bundle of fibers and continue to tease apart until single fiber segments are isolated from the bundle.
   5. Collect a minimum of 50 fibers that are at least ~1 mm in length.
   6. Move the fiber to an empty space on the Petri dish. Inspect single-fiber segments under higher magnification (x 50). Ensure it is a single fiber by further separating the fiber at the end. If the fiber breaks instead of separating, it is a single fiber.
3. Denaturation of fibers
   1. Using forceps, gently collect the fiber and place it directly into the aliquoted denaturing buffer (do not let the forceps encounter the buffer).
   2. Check the forceps under the microscope to ensure the fiber has been successfully removed. Close the tube and tap the bottom of the tube firmly on the bench three times to ensure that the fiber moves into the buffer.
   3. Wipe the forceps clean using lint-free tissue before moving on to the next fiber. Repeat this process until all fibers are collected.
   4. Vortex the fiber samples and briefly centrifuge for 5 s at 2,500 × g to draw the sample to the bottom of the tube.
      NOTE: The centrifugation duration (5 s) is timed from the start (0 × g) until the speed reaches ~2,500 × g.
   5. Leave the samples at room temperature for 1 h. Store at -80 °C for future use. Freeze and then thaw the samples before use.

3. Dot blotting

1. Membrane preparation and activation
   1. Measure, label, and cut to size one polyvinylidene difluoride (PVDF) membrane (0.2 µm pore size) to fit 50 samples (~10.5 cm x 5.5 cm).
   2. Mark the left border numerically from top to bottom (1 to 10) and the top border alphabetically from left to right (A to E) spaced 1 cm apart.
   3. Prepare four sheets of filter paper with the dimensions of 12.5 cm x 7.5 cm.
   4. Pile two sheets together to form a stack. Presoak the filter paper stack in 1x transfer buffer.
   5. Place the membrane in a container. Pour 95% ethanol over the membrane, with enough volume to entirely immerse the membrane (10-15 mL). Agitate on the rocker for 1 min.
   6. Recollect the ethanol, immerse the membrane in 1x transfer buffer (~15 mL), and rock for a further 2 min.
      NOTE: Both ethanol and transfer buffer can be reused for future dot blotting experiments until sedimentation forms (change after six uses).
   7. Lay the transfer buffer-soaked filter paper stack on a flat movable surface such as a large lid and flatten with the roller. Position the PVDF membrane on the stack using a gel releaser or tweezers.
   8. Place a single sheet of dry filter paper on top of the membrane and move the roller over the filter paper to soak up any excess buffer on the membrane surface. Remove the top filter paper without rubbing the membrane.
      CAUTION: Methanol (Class 3, sub-risk 6.1) and ethanol (Class 3) are hazardous. Refer to the material safety datasheet (MSDS) for safe handling recommendations.
2. Sample absorption to the membrane
   1. Thaw the fiber samples, perform a brief centrifuge (5 s) at room temperature as in step 2.3.4 and mix the sample thoroughly.
   2. Without touching the membrane with the pipette tip, slowly deposit a ~1 µL drop of each sample onto the membrane in the designated area. The dimensions of the designated area per fiber sample are ~1 cm x 1 cm. Aim to spot the sample in the center of this area.
   3. Allow the sample droplets to soak through the membrane for at least 15 min. Ensure no sample buffer remains and is completely absorbed.
   4. Using plastic tweezers, carefully lift the membrane off the damp filter paper stack, place it on a single dry sheet of filter paper, and dehydrate the membrane for at least 5 min.
   5. Once the sample spots turn completely white, reactivate the membrane.
3. Membrane reactivation and block
   1. Reactivate the membrane by repeating steps 3.1.5 and 3.1.6.
   2. Place the membrane in wash buffer and rinse for 5 min with rocking. Discard the wash buffer.
   3. Incubate the membrane in blocking buffer for 30 min while rocking.
   4. Rinse the membrane 3x with wash buffer until the buffer is no longer cloudy.
   5. Leave the membrane in the final wash on the rocker.

4. Immunolabeling

1. Detection of MHCIIa
   1. Dilute the MHCIIa primary antibody at 1 in 200 in 10 mL of Bovine Serum Albumin (BSA) buffer.
   2. Discard the wash buffer from the membrane and then pour the diluted MHCIIa antibody onto the membrane.
   3. Place the container (with the membrane in MHCIIa) on the rocker at room temperature for 2 h or at 4 °C overnight.
   4. Recollect the MHCIIa antibody and store it at 4 °C. Wash the membrane with blocking buffer for 2 min on the rocker.
      NOTE: All primary antibodies can be reused up to five times as long as the buffer remains clear.
   5. Rinse twice more; 5 min each time; three washes in total.
   6. Dilute mouse Immunoglobulin G Horseradish Peroxidase (IgG-HRP) secondary antibody at 1 in 20,000 in 10 mL of blocking buffer and add to the membrane.
   7. Discard the wash buffer and incubate the membrane in mouse IgG-HRP secondary with rocking for 1 h.
   8. Discard the secondary and wash the membrane in wash buffer (2, 5, 5 min).
   9. Leave the membrane soaking in the final wash until ready for imaging.
   10. Turn on the gel imager, wait 15 min for the imager to reach the required operating temperature, and then open the imaging software (see Table of Materials).
   11. In the software window, click New Single Channel Protocol (Figure 3A). For a white light image of the membrane, under Applications | Blots | click Colorimetric (Figure 3B).
   12. Click on the gel area; each option is programmed for specific dimensions to fit the size of various gel types. Select the appropriate option to best fit the dimensions of the membrane (Figure 3C).
   13. Prepare the ECL by combining luminol and peroxidase reagents at a 1:1 ratio. Make a sufficient volume of ECL mixture to cover the entire membrane, typically a total volume of 800-1,000 µL is required.
   14. Place the membrane on a large clear plastic tray and disperse the ECL mixture onto the membrane.
   15. Place the membrane onto the imaging plate.
   16. Click on Position Gel (yellow button) for a live camera view. Click hold and drag the zoom button to maximize the imaging area of the membrane. At this point, straighten the position of the membrane if required. 
   17. Click Run Protocol (green button) and wait for a colorimetric image of the membrane to be produced. Save this image to assist in matching the dots to the corresponding fiber sample.
   18. Without moving the membrane, change the application on the software by clicking the option for a 2 x 2 binning (Chemi High Resolution) (Figure 3D).
   19. Under Image exposure and software, optimize the exposure time, click Faint Bands (Figure 3E) | Signal Accumulation Mode.
   20. Under Setup, type in 1 s for the first image, 30 s for the last image, and 30 total images (Figure 3F).
   21. Click Run Protocol.
   22. Save an image at the following stages: before signal saturation (all visible dots are black), initial signal saturation (some dots begin to saturate), and oversaturated spots (all spots with strong signal detected are saturated).
   23. Save all required images before ending signal accumulation. Take the membrane out of the imager.
   24. Using the gel releaser, place the membrane back into the container with the wash buffer.
2. Detection of MHCI
   1. Pour the wash buffer out and treat the membrane with stripping buffer for 30 min at 37 °C (ensure the membrane is completely immersed).
   2. Recollect the stripping buffer and rinse the membrane in wash buffer for 5 min with rocking.
      NOTE: Stripping buffer can be reused 10x before discarding.
   3. Pour out the wash buffer and this time apply MHCI primary antibody diluted at a starting concentration of 1 in 200 (range: 1 in 200 to 1 in 500). Rock the membrane at room temperature for 1 h.
   4. After incubating in MHCI primary antibody, recollect the antibody and wash the membrane as in steps 4.1.4 and 4.1.5.
   5. Dilute mouse IgM HRP secondary antibody in blocking buffer at 1 in 20,000. Pour out the blocking buffer from the membrane and incubate in the mouse IgM secondary as in step 4.1.7.
   6. Wash and image capture detected MHCI as in steps 4.1.8 to 4.1.24.
      CAUTION: Stripping buffer contains Organo Phosphine 3-5% (Class 8). Refer to the MSDS for safe handling recommendations.
3. Detection of potential MHCIIx using Actin
   1. Strip the membrane once more (steps 4.2.1 and 4.2.2).
   2. Repeat section 4.1, this time applying Actin primary antibody diluted at 1 in 500 in BSA buffer and then rabbit HRP secondary antibody diluted at 1 in 20,000 in blocking buffer.

5. Fiber type identification

1. MHCI, MHCIIa, and Actin signal analysis
   1. Become familiar with the signal intensity panel (Figure 4A). Make a note of steps 5.1.2 to 5.1.4.
   2. A saturated signal intensity qualitatively indicates that the protein detection is strong.
   3. A moderate indicates that the target protein is present at a medium level.
   4. A faint signal indicates little target protein is present.
   5. Use the signal intensity panel to categorize fibers with 'saturated,' 'moderate,' or 'faint' signal intensity (Figure 4A).
   6. Carry out fiber type identification by comparing MHCIIa and MHCI results first (Figure 4B, left and middle blots).
   7. Record fibers with saturated or moderate signal intensity of only one MHC isoform.
   8. If the MHC isoform signal is 'faint,' leave the fiber unmarked (see Figure 4B).
   9. Lastly, where Actin is observed (moderate or saturated signal intensity) in the fibers with no MHCI or IIa detected, record it as a potential type IIx fiber (Figure 4B, right blot).
   10. Record fibers with faint MHC isoform signal but Actin detected (moderate or saturated intensity) as unidentified.
   11. Discard samples with faint or no detection (no fiber collected) for all three target proteins (Figure 4C).
   12. Record any fiber where both MHCI and MHCII signals are detected with a 'saturated' or moderate signal. These samples are not for use in fiber type-specific preparation.
2. Preparation of fiber type-specific samples
   1. Prepare a separate type I and type II sample for each biopsy. For a type II sample, combine the minimum required number of fibers (calculated in step 2.1.5) in which MHCIIa is detected at saturated or moderate levels.
   2. In a separate tube labeled as type I, repeat this process for fibers in which MHCI is detected (Figure 4D).
   3. Use fibers with moderate signal intensity if there are not enough fibers with saturated signals.
   4. Combine any potential IIx fibers.
   5. Immediately use fiber type-specific samples for western blotting or store them at -80 °C for future use.

6. Western blot fiber type confirmation

1. Utilize 10 µL of each fiber type-specific sample, (the minimum sample volume required, see step 2.1.5).
2. Separate samples using the specified precast gel via SDS-PAGE according to the manufacturer's protocol.
3. Use a gel imager to detect the protein in the gel according to the manufacturer's protocol.
4. Place the gel into cold 1x transfer buffer for 10 min.
5. Wet transfer the proteins onto a 0.45 µm nitrocellulose membrane (9.5 cm x 13.5 cm) according to the manufacturer's protocol.
6. After transfer, briefly rinse the membrane with ultrapure H₂O in a container. Pour out the ultrapure water.
7. Add 10 mL of antibody signal enhancer solution to the membrane and rock at room temperature for 10 min.
8. Recollect the antibody signal enhancer solution and wash 5x (5 s per wash) with ultrapure H₂O.
   NOTE: Antibody signal enhancer solution can be used 5x before discarding.
9. Pour out the last wash, add blocking solution, and rock at room temperature for 1 h.
10. Use a clean scalpel blade or scissors to cut horizontally across the membrane at a molecular weight that ensures that proteins that are 180 kDa and above are on the top section of the membrane.
11. Use this top section for further fiber type confirmation.
12. Use the portion below 180 kDa to detect different target proteins.
13. On the top section of the membrane, follow section 4.1 with MHCIIx primary and mouse Immunoglobulin M (IgM) HRP secondary antibody.
14. Strip the membrane (as described in sections 4.2.1 and 4.2.2) before repeating the process with MHCIIa IgG primary and mouse IgG HRP secondary antibody.
15. Strip once more and finally apply MHCI IgM primary and mouse IgM HRP secondary antibody.
    NOTE: This step is qualitative and so any protein loss due to membrane stripping is not a concern.
16. Compare the signal detected across the different MHC isoforms (as seen in Figure 5B). When the signal intensity is detected at moderate to saturated levels in the expected sample (MHCI - type I, MHCIIa - type II and MHCIIx - type IIx) the fiber type-specific samples will be recorded as reliable for use in future studies.
17. Record the sample as unreliable when more than one MHC isoform is equally detected in a sample.
    CAUTION: Antibody enhancer solution contains sodium hydroxide 50%. Refer to MSDS for safe handling recommendations.

Results

Identification of individual MHCI, MHCIIa, and MHCIIx muscle fibers using dot blotting
A feature of MyDoBID is the categorization of the varying MHC and Actin signal intensity strength in a given fiber (Figure 4A). Fiber type was identified by the presence or absence of MHCI and IIa isoforms (Figure 4B). Six fibers showed no MHC or actin detection, indicating that there was no fiber collected. The results of this specific dot blot were the i...

Discussion

Fiber collection
Based on several years of experience, most researchers can master this technique; however, practice leads to faster and more efficient fiber collection for downstream analyses. To be able to isolate 30 single fiber segments of a quality for pooling, it is recommended that 50 fiber segments are collected per sample. Studying the fiber collection video carefully and after performing two practice sessions (~50 fibers per session) is recommended to achieve a reasonable standard. All fi...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1x Denaturing buffer	Make according to recipe	Constituents can be sourced from Sigma-Aldrich or other chemical distributing companies	1x denaturing buffer is made by diluting 3x denaturing buffer 1 in 3 v/v with 1X Tris-HCl (pH 6.8). Store at -20 °C.
3x Denaturing buffer	Make according to recipe	Constituents can be sourced from Sigma-Aldrich or other chemical distributors	3x denaturing buffer contains: 0.125M Tris-HCI, 10% glycerol, 4% SDS, 4 M urea, 10% 2-mercaptoethanol, and 0.001% bromophenol blue, pH 6.8.  Store at -20 °C.
95% Ethanol	N/A	100% ethanol can be sourced from any company	Diluted to 95% with ultra-pure H2O.
Actin rabbit polyclonal antibody	Sigma-Aldrich	A2066	Dilute 1 in 1,000 with BSA buffer.
Analytical scales	Mettler Toledo	Model number: MSZ042101
Antibody enhancer	Thermo Fischer Scientific	32110	Product name is Miser Antibody Extender Solution NC.
Beaker (100 mL)	N/A	N/A
Benchtop centrifuge	Eppendorf	5452	Model name: Mini Spin.
Blocking buffer: 5% Skim milk in Wash buffer.	Diploma	Store bought
BSA buffer: 1 % BSA/PBST, 0.02 % NaN3	BSA: Sigma-Aldrich              PBS: Bio-Rad Laboratories. NaN3 : Sigma-Aldrich	BSA: A6003-25G                         10x PBS: 1610780                       NaN3: S2002	Bovine serum albumin (BSA), Phosphate-buffered saline (PBS), and Sodium azide (NaN3). Store at 4 °C.
Cassette opening lever	Bio-Rad Laboratories	4560000	Used to open the precast gel cassettes.
Chemidoc MP Imager	Bio-Rad Laboratories	Model number: Universal hood III	Any imaging system with Stain-Free gel imaging capabilities.
Criterion blotter	Bio-Rad Laboratories	1704070	Includes ice pack, transfer tray, roller, 2 cassette holders, filter paper, foam pads and lid with cables.
Criterion Cell (Bio-Rad)	Bio-Rad Laboratories	1656001
ECL (enhanced chemiluminescence)	Bio-Rad Laboratories	1705062	Product name: Clarity Max Western ECL Substrate.
Electrophoresis buffer 1x Tris Glycine SDS (TGS)	Bio-Rad Laboratories	1610772	Dilute 10x TGS 1 in 10 with ultra-pure H2O.
Filter paper, 0.34 mm thick	Whatmann	3030917	Bulk size 3 MM, pack 100 sheets, 315 x 335 mm.
Fine tissue dissecting forceps	Dumont	F6521-1EA	Jeweller’s forceps no. 5.
Flat plastic tray/lid	N/A	N/A	Large enough to place the membrane on. Ensure the surface is completely flat.
Freeze-drying System	Labconco	7750030	Freezone 4.5 L with glass chamber sample stage.
Freezer -80 oC	N/A	N/A	Any freezer with a constant temperature of -80 °C is suitable.
Gel releasers	1653320	Bio-Rad	Slide under the membrane to gather or move the membrane.
Grey lead pencil	N/A	N/A
Image lab software	Bio-Rad Laboratories	N/A	Figures refers to software version 5.2.1 but other versions can used.
Incubator	Bio-Rad Laboratories	1660521	Any incubator that can be set to 37 °C would suffice.
Lamp	N/A	N/A
Magnetic stirrer with flea	N/A	N/A
Membrane roller	Bio-Rad Laboratories	1651279	Can be purchased in the Transfer bundle pack. However, if this product is not available, any smooth surface cylindrical tube long enough to roll over the membrane would suffice.
Microcentrifuge tubes  (0.6 mL)	N/A	N/A
Mouse IgG HRP secondary	Thermo Fisher Scientific	31430	Goat anti-Mouse IgG (H+L), RRID AB_228341. Dilute at 1 in 20,000 in blocking buffer.
Mouse IgM HRP secondary	Abcam	ab97230	Goat Anti-Mouse IgM mu chain. Use at the same dilution as mouse IgG.
Myosin Heavy Chain I (MHCI) primary antibody	DSHB	A4.840	Dilution range: 1 in 200 to 1 in 500 in BSA buffer.
Myosin Heavy Chain IIa (MHCIIa) primary antibody	DSHB	A4.74	Dilution range: 1 in 200 to 1 in 500 in BSA buffer.
Myosin Heavy Chain IIx (MHCIIx) primary antibody	DSHB	6H1	Dilution range: 1 in 200 to 1 in 500 in BSA buffer.
Nitrocellulose Membrane 0.45 µm	Bio-Rad Laboratories	1620115	For Western blotting.
Petri dish lid	N/A	N/A
Plastic tweezers	N/A	N/A
Power Pack	Bio-Rad Laboratories	164-5050	Product name: Basic power supply.
Protein ladder	Thermo Fisher Scientific	26616	PageRuler Prestained Protein Ladder, 10 to 180 kDa.
PVDF Membrane 0.2 µm	Bio-Rad Laboratories	1620177
Rabbit HRP secondary	Thermo Fisher Scientific	31460	Goat anti-Rabbit IgG (H+L), RRID AB_228341. Dilution same as mouse secondary antibodies.
Rocker	N/A	N/A
Ruler	N/A	N/A
Scissors	N/A	N/A
Stereomicroscope	Motic	SMZ-168
Stripping buffer	Thermo Fisher Scientific	21059	Product name: Restore Western Blot Stripping Buffer.
Tissue (lint free)	Kimberly-Clark professional	34120	Product name: Kimwipe.
Transfer buffer (1x Tris Glycine buffer (TG), 20% Methanol)	TG: Bio-Rad Laboratories   Methanol: Merck	TG buffer: 1610771           Methanol: 1.06018	dilute 10x TG buffer with ultra-pure H2O to 1x. Add 100% Methanol to a final concentration of 20% Methanol. Store at 4 °C.
Transfer tray	Bio-Rad Laboratories	1704089
UV-activation precast gel	Bio-Rad Laboratories	5678085	Gel type: 4–15% Criterion TGX Stain-Free Protein Gel, 26 well, 15 µL.
Vortex	N/A	N/A
Wash buffer (1x TBST)	10x TBS: Astral Scientific  Tween 20: Sigma	BIOA0027-4L	1x TBST recipe: 10x Tris-buffered saline (TBS) is diluted down to 1x with ultra-pure H2O, Tween 20 is added to a final concentration of 0.1%. Store buffer at 4 °C.
Wash containers	Sistema	Store bought	Any tupperware container, that suits the approximate dimensions of the membrane would suffice.