MultiBac System-Based Purification and Biophysical Characterization of Human Myosin-7a

NOTE: Here we describe a protocol for synthesizing the intact human myosin-7a holoenzyme and characterizing its motility in vitro. This protocol is divided into three sections: first, expressing the human myosin-7a using the MultiBac baculovirus protein expression system; Second, purifying myosin-7a light chains separately using the E.coli His6-SUMO system; and lastly, studying the motility of human myosin-7a using the actin-filament gliding assay.

1. MultiBac system-based myosin-7a complex production

1. Creating baculovirus multigene cassette for expressing human myosin-7a complex (16 days)
   NOTE: It requires one cloning cycle (4 days) to insert the cDNAs of myosin-7a subunits into the pACEBac1 vectors and three cloning cycles to complete the step-wise ligation of all multiplication modules required for expressing the myosin-7a complex, resulting in a total of 16 days.
   1. Clone the cDNAs encoding the myosin-7a heavy chain (HC), calmodulin, CALML4, and RLC into the pACEBac1 vectors using a commercial kit following the manufacturer's protocols (see Table of Materials; Figure 1A). A FLAG tag is inserted at the C-terminus of the myosin-7a HC to aid in purification.
      NOTE: Mammalian myosin-7a is produced as two splicing isoforms, differing only by a segment of 11 amino acids at the N-terminus²³. This short N-terminal extension plays a critical role in regulating the ATPase activity of myosin-7a¹⁴. It has been shown that adding an N-terminal tag can disrupt the normal regulation by this N-terminal extension and significantly reduce the motor's enzymatic activity and motility¹⁴. Therefore, a C-terminal FLAG tag is employed to avoid disrupting the protein's natural regulation.
   2. Construct the multigene expression cassette for the myosin-7a holoenzyme using homing endonuclease/BstXI multiplication as described below (Figure 1B).
      NOTE: The homing endonuclease I-CeuI produces cohesive ends that are compatible with the ends generated by the BstXI digest. Upon ligation, a homing endonuclease/BstXI hybrid restriction site is created that cannot be cut by either enzyme. The same procedure can be repeated to integrate more multiplication modules. If multiple restriction sites for I-CeuI and BstXI are present on the insert or vector constructs, these additional sites must be eliminated by site-directed mutagenesis to ensure that the I-CeuI and BstXI cutting sites are unique.
   3. Insert preparation: Digest the insert construct (e.g., pACEBac-M7a HC) with the homing endonuclease I-CeuI (see Table of Materials), and then purify the linearized plasmid using a PCR Purification Kit (see Table of Materials). Further, digest the linearized plasmid with BstXI (see Table of Materials), separate the fragment containing the gene expression cassette using gel electrophoresis, and purify using a Gel Extraction Kit (see Table of Materials).
      NOTE: I-CeuI and BstXI require different buffer conditions, according to manufacturer's protocols, to achieve 100% activity. In addition, a complete cut with I-CeuI requires a minimum of 3 h, while BstXI requires only 15 min of incubation time. The digestions thus need to be performed sequentially.
   4. Vector preparation: Linearize the vector construct (e.g., pACEBac-CaM) through BstXI digestion. Alkaline Phosphatase is included in the vector preparation to prevent the vector plasmid from ligating to itself (see Table of Materials) . Obtain the digestion product by electrophoresis and purify it with a Gel Extraction Kit.
   5. Ligation: Ligate the I-CeuI/BstXI-treated insert and the BstXI-treated vector with T4 DNA Ligase (see Table of Materials). Perform ligation reactions at room temperature (20 °C -25 °C) for 10 min with a 1:1 ratio by mass of the insert DNA to vector DNA, keeping the total mass close to 100 ng in each ligation reaction.
   6. Transformation and plasmid analysis: Transform the ligation mixtures into DH5α cells (see Table of Materials) following the manufacturer's recommendations and screen the positive colonies for gentamicin resistance. Purify the plasmids using a commercial kit (see Table of Materials), and verify the correct clones (e.g., plasmids containing both M7a HC and CaM) based on specific restriction digestion patterns and DNA sequencing of the inserts.
   7. Integration of multiple gene expression cassettes: Repeat steps 1.1.2 through 1.1.6 to integrate additional gene cassettes expressing other myosin-7a subunits.
2. Recombinant bacmid DNA production and purification (4 days; Figure 1C).
   NOTE: Day 1 – Bacterial transformation and growing of transformed colonies. Days 2-3 – Growing colonies and screening for successful transformation. Day 3 – Selecting and streaking positive colonies, starting overnight inoculation. Day 4 – Purification of bacmid DNA.
   1. Transform DH10Bac competent cells (see Table of Materials) with 1 ng of the sequenced pACE-M7aHC-RLC-CALML4-CaM plasmid following the manufacturer's recommendations.
   2. Plate 20-200 µL of the cells on an agar plate containing 50 µg/mL kanamycin, 7 µg/mL gentamicin, 10 µg/mL tetracycline, 100 µg/mL halogenated indolyl-β-galactoside, and 40 µg/mL IPTG (see Table of Materials) and incubate the plate at 37˚C for 48 h to allow for the development of deep blue color in negative clones.
      NOTE: The halogenated indolyl-β-galactoside stains colonies that continue to express LacZ (indicating the transposition was unsuccessful), enabling the selection of white colonies where transposition was successful.
   3. Carefully select an isolated white colony and grow cells overnight in 5 mL of LB medium containing 50 µg/mL kanamycin, 7 µg/mL gentamicin, and 10 µg/mL tetracycline at 37 ˚C in an orbital shaker at 225 rpm.
   4. Centrifuge the culture at 2,465 x g for 10 min to pellet the E coli. Discard the supernatant and resuspend the pellet in 300 µL of Buffer P1 from the Kit.
      NOTE: While buffers from the Miniprep kit are used here, the protocol for purifying the bacmid product varies from the protocol provided with the kit.
   5. Add 300 µL of the Buffer P2 from the Kit. Mix by inverting the tubes a few times, and then incubate at room temperature for 3 min.
   6. Add 300 µL of the N3 buffer from the Kit and mix by inversion to stop the lysis reaction. Centrifuge at 17,885 x g for 10 min at 4 °C.
   7. Transfer the supernatant to a clean sterile tube and repeat the centrifugation for another 10 min.
   8. Transfer 800 µL of the supernatant to another sterile 1.7 mL microcentrifuge tube (see Table of Materials) and add 600 µL of cold isopropanol (see Table of Materials), mixing by rigorous inversion. Then centrifuge at 17,885 x g for 20 min at 4 °C.
   9. Discard the supernatant and locate the small, white pellet. Wash the pellet with 800 µL of cold 70% ethanol (see Table of Materials) and centrifuge at 17,885 x g for 5 min at 4 °C. Carefully add ethanol along the tube wall to avoid disturbing the pellet.
   10. Discard the supernatant and repeat the ethanol wash once. Air dry the pellet at room temperature for 5 min. Carefully aspirate any excess liquid, if necessary.
   11. Dissolve the pellet with 20 µL of EB buffer from the Kit. Ensure the pellet is fully dissolved by flicking the tube or gently mixing by pipetting. Determine concentration using a spectrophotometer (see Table of Materials).
   12. Adjust the buffer volume as needed to make the bacmid concentration at approximately 1 µg/µL. Store bacmid DNA at 4 °C until ready for P0 virus production.
       NOTE: The purified bacmid can be kept at 4 ˚C and remains effective for several months. We have seen successful transfections with bacmid products up to 1 year old. Alternatively, the bacmid can be aliquoted and stored at -20 ˚C. Multiple freeze/thaw cycles should be avoided as they decrease the transfection efficiency.
3. Transfect Sf9 insect cells with bacmid to produce recombinant baculovirus (6 days; Figure 1D)
   ​NOTE: Day 1 – Transfection of cells with bacmid DNA. Days 2-6 – Observe growth and expression of cells. Day 6 – Collection of P0 virus.
   1. Add Sf9 cells in culture media (see Table of Materials) to a 6-well plate at a concentration of 0.33 x 10⁶ cells/mL with 3 mL per well. Let the plate sit at room temperature for 30 min to allow the cells to attach to the bottom of the plate.
   2. For each well, prepare a transfection mixture by combining 10 µL of cationic lipid transfection reagent (see Table of Materials) with 250 µL of improved Minimal Essential Medium (MEM) media (see Table of Materials) in a sterile microcentrifuge tube. Mix by inversion and incubate at room temperature for 5 min.
   3. Add 1 µg (based on concentration determined in step 1.2.12) of undiluted myosin-7a bacmid to the transfection mixture. Mix by inversion and incubate at room temperature for 5 min.
   4. Equally distribute the entire DNA-lipid mixture dropwise into the wells. Leave one well for non-transfected cells as a negative control.
   5. Seal the 6-well plate with film (see Table of Materials) and incubate at 27 °C for 6 days. Observe growth and detachment throughout this time. If there is a fluorescent tag present on the construct, check for fluorescence development (Figure 2).
   6. When the transfected cells show signs of late-stage infection, transfer the medium containing the virus from infected wells to a 15 mL conical tube and centrifuge at 805 x g for 10 min.
   7. Transfer the supernatant to a clean 15 mL conical tube, wrap it in aluminum foil, and store it at 4 °C. This product is now the P0 virus and can be stored and used effectively for up to four months from our experience.
4. Amplify baculovirus stock (3-5 days ; Figure 1E)
   1. Prepare 100 mL (or desired volume) of Sf9 cells at a concentration of 2 x 10⁶ cells/mL in Sf9 cell culture media in a 250 mL Erlenmeyer flask (see Table of Materials).
   2. Add 1: 100 ratio (v/v) of P0 virus stock to the Sf9 cell suspension culture and incubate at 27 ˚C in an orbital shaker at 135 rpm.
   3. Monitor growth every 24 h until cells achieve ~90% cell death. Once at this range, centrifuge the cell suspension at 500 x g for 10 min and collect the supernatant containing the P1 virus.
   4. Filter the supernatant using 0.22 µm vacuum filtration (see Table of Materials). Keep the filtered product in the dark and store it at 4 °C. This product is now the P1 virus.
      NOTE: Virus titer decreases with long-term storage at 4 ˚C. Consider making fresh virus stock if it has been stored for more than 4 months, or titer the virus stock before use in expression³⁴.
5. Protein Expression (60 -72 h between beginning the expression and collection of the cell pellet ; Figure 1E)
   1. Prepare log-phase growing Sf9 cells at a concentration of 2 x 10⁶ cells/mL and add P1 virus at a ratio of 12 mL of virus to 300 mL of cell suspension.
      NOTE: The protein yield is approximately 1 mg/L. It is recommended to use a minimum of 1.5 L of cell suspension for this protein expression.
   2. Culture the cells at 27 °C in an orbital shaker at 135 rpm for approximately 60 h.
      NOTE: Small samples can be collected at different time points and analyzed using SDS gels to assess protein expression levels. Additionally, if the proteins are fused with fluorescent tags, expression levels can be evaluated using fluorescence microscopy. The cells should be harvested no later than the onset of cell death.
   3. Centrifuge the cell suspension at 3,735 x g for 5 min at 4 °C to pellet the cells. The pellets can be stored at -80 ˚C until further use or preserved for long-term storage. We typically use cell pellets within weeks but have recovered active protein several months after freezing.
6. Protein purification (2 days ; Figure 1F)
   NOTE: Day 1 – Protein extraction and purification using FLAG-affinity chromatography. Day 2 – Aliquot and freeze samples after overnight dialysis. The protocol described below is for purifying myosin-7a from 1.5 L cell pellets.
   1. Prepare Master Buffer containing 10 mM MOPS (pH 7.2), 500 mM NaCl, 10 mM MgCl₂, 1 mM EGTA, 1 µg/mL leupeptin, and 100 µg/mL phenylmethylsulfonyl fluoride (PMSF). Filter it with a 0.22 µm pore size filter and store the buffer at 4 ˚C (see Table of Materials).
      NOTE: PMSF is unstable in aqueous solutions. Prepare the PMSF stock solution as 1 M in 100% ethanol and store it at -20 ˚C for up to 6 months.
   2. Prepare 75 mL of Extraction Buffer by supplementing the Master Buffer with 3 EDTA-free protease inhibitor tablets, 2 mM ATP, and 0.1 mM dithiothreitol (DTT; see Table of Materials).
   3. Add 75 mL of Extraction Buffer to the pellet while it is still frozen. Leave the pellet in the Extraction Buffer on ice until it thaws. Use a serological pipette to break up the pellet to accelerate the thawing process.
   4. Using a ½-inch (13 mm) tip, sonicate the resuspension on ice for 10 min at 50% amplitude, 5 s on, 5 s off.
   5. Centrifuge the total lysate at 33,746 x g for 30 min at 4 °C. During the centrifugation, prepare 1.5 mL of FLAG resin by washing 3 mL of the 50% slurry of Anti-FLAG affinity resin (see Table of Materials) with 40 mL of PBS. Pellet the resin by centrifuging at 800 x g for 2 min at 4 °C. Carefully remove the PBS without disturbing the resin.
   6. Add the supernatant from the lysate centrifugation to the washed resin and incubate with continuous rotation at 4 °C for 1 h.
   7. Pellet the resin by centrifuging at 800 x g for 2 min at 4°C. Without disturbing the resin, remove the supernatant.
   8. Resuspend the resin in 40 mL of Master Buffer and centrifuge at 800 x g for 2 min at 4 °C. Carefully remove the supernatant without disturbing the resin. Repeat the wash 1x.
   9. Transfer the washed resin to two polypropylene spin columns (see Table of Materials). Wash each column 1x with 2 mL of Master Buffer. Remove the Master Buffer by centrifuging the column at 1,400 x g for 2 min at 4 °C. The resin will be packed on the bottom.
   10. Prepare an Elution Buffer by adding 100 µg/mL of FLAG-peptide (see Table of Materials) to the Master Buffer.
   11. Add 300 µL of Elution Buffer to the packed resin in each column and gently mix by pipetting to ensure the resin is completely hydrated by the Elution Buffer. Let the resin incubate with the Elution Buffer on ice for 10 min.
   12. Centrifuge the columns at 1,400 x g for 2 min at 4 °C. Transfer the flowthrough to a clean microcentrifuge tube and keep it on ice. This is the first fraction.
   13. Repeat steps 1.6.11-1.6.12 such that 4 fractions are collected from each column.
   14. Perform SDS-PAGE gel electrophoresis³⁵ with the elution fractions to determine their relative concentrations. While the gel runs, prepare a Dialysis Buffer containing 500 mM NaCl, 10 mM MOPS, 2 mM MgCl₂, 0.1 mM EGTA, and 1 mM DTT.
   15. Combine the most concentrated fractions. Transfer the sample to a 10K MWCO dialysis cassette (see Table of Materials) and dialyze overnight at 4 °C. Use 1 L dialysis buffer for approximately 500 µL of protein. Perform at least three changes of the dialysis buffer.
       NOTE: The spin column and centrifugation elution method allow proteins to be eluted at high concentrations (0.5-1 mg/mL). Further concentrating steps are not usually required. However, if more concentrated proteins are desired, load samples onto 100,000 MWCO concentrating tubes (see Table of Materials) and centrifuge at 1,400 x g for 5 min at 4 ˚C. Repeat this process until the desired protein solution volume is achieved.
   16. The next day, carefully unload the sample from the dialysis cassette. Aliquot 15 µL per PCR tube and drop the tubes into a container of liquid nitrogen for flash-freezing. The frozen proteins can be stored at -80 ˚C for future use (Figure 3A).
       NOTE: The yield of myosin-7a using this method is usually between 0.5 and 1 mg/L.

2. Purifying the light chains RLC and CALML4 using the E. coli His6-SUMO system (7 days )

NOTE: Days 1-4 – Cloning and purification of the plasmids. Day 5 – Bacterial transformation. Days 6-7 – Purification, aliquoting, and freezing of final proteins.

1. Clone the cDNAs encoding RLC and CALML4 into a pET-SUMO vector, which contains a His6 sequence prior to SUMO to aid in purification. When needed, the His6-SUMO domain can be cleaved from the fusion protein using a SENP2 protease^26,36.
2. For each light chain protein, transform BL21 E. coli competent cells (see Table of Materials) with the plasmid and start a 5 mL overnight liquid culture with the transformed cells.
3. The next day, inoculate the 5 mL overnight culture into 1 L of Luria-Bertani (LB) broth containing 50 µg/mL kanamycin. Allow the culture to grow at 37 ˚C with continuous shaking at 270 rpm until the optical density (O.D.) reaches 0.6 – 1.0.
   NOTE: The O.D. value doubles approximately every 20 min once it reaches 0.2. Monitor the O.D. frequently.
4. Initiate protein expression by adding 250 µM isopropyl b-D-1-thiogalactopyranoside (IPTG) to the culture. Incubate for 3-5 h at 37 ˚C or 16 ˚C overnight with continuous shaking at 270 rpm.
5. Pellet the E. coli by centrifuging the cell suspension at 4,347 x g for 15 min at 4 °C. Discard the supernatant.
6. Prepare Resuspension Buffer containing 50 mM Tris-HCl, 1 M NaCl, 5 mM Imidazole, and 10% glycerol, pH 7.4 (see Table of Materials). Keep on ice.
7. Resuspend the pellet in 15-25 mL of Resuspension Buffer and transfer the suspension to a clean beaker, adjusting the total volume to 35 mL. Add DNase and RNase to the suspension at a 1:1000 (v/v) ratio for a final concentration of 1 µg/mL and 2 µg/mL, respectively. Add one EDTA-free protease inhibitor tablet, 4 mM 2-mercaptoethanol, and 1% Triton X to the suspension (see Table of Materials). Keep on ice for 20 min.
8. Sonicate the pellet suspension in ice water with 1 s on, and 1 s off for a total of 1 min at 100% amplitude. Leave the homogenized lysate at 4 ˚C with continuous rotation for 2 h.
   NOTE: The addition of Triton X aids in lysing the cells in preparation for sonication.
9. Centrifuge the lysates at 26,900 x g for 45 min at 4 ˚C. While this is occurring, begin preparing the cobalt resin (see Table of Materials). Use 500 µL of cobalt resin for 1 L of E. coli culture. Wash the resin 2x with ultrapure water and once with Suspension Buffer by centrifuging at 4,347 x g for 2 min at 4 °C.
10. Following lysate centrifugation, combine the supernatant with the washed resin and rock gently at 4 ˚C for 30 min.
11. Centrifuge the lysate-resin suspension at 4,347 x g for 2 min at 4 °C. The resin will be packed at the bottom of the tube. Without disturbing the resin, remove the supernatant.
12. Wash the resin with Resuspension Buffer by centrifuging at 4,347 x g for 2 min at 4 °C. Remove the supernatant. Repeat washes until the supernatant is colorless.
13. Prepare Elution Buffer containing 50 mM Tris HCl, 300 mM NaCl, 5 mM Imidazole, 10% glycerol, 4 mM 2-mercaptoethanol, and 0.25 µM SENP2 protease (see Table of Materials). Keep it on ice.
14. Resuspend the resin in 1 mL of Resuspension Buffer and incubate it overnight at 4 °C with continuous rotation.
15. The next day, centrifuge at 4,347 x g for 2 min to pellet the resin. Collect the supernatant, which contains the cleaved light chain protein and the protease.
16. Add 1 mL of Elution Buffer to the resin. Repeat centrifugation and collect the supernatant.
17. Remove the protease by fast protein liquid chromatography (FPLC). Briefly, prepare the size exclusion column (see Table of Materials) with a buffer containing 50 mM Tris-HCl, 300 mM NaCl, 10% glycerol, and 5 mM DTT (pH 7.4). Concentrate the supernatant to 500 µL using a 10,000 MWCO concentrating tube. Load the concentrated sample onto a column connected to an automated chromatography system using a syringe, flow through with a buffer containing 50 mM Tris-HCl, 300 mM NaCl, 10% glycerol, and 5 mM DTT (pH 7.4). Monitor the collected fractions using an ultraviolet spectrophotometry. Collect the protein fractions with a molecular weight corresponding to the light chains.
    NOTE: The protease can also be removed by affinity chromatography if a purification tag (e.g., GST tag) is engineered into the protease.
18. Run an SDS-PAGE gel with the elution fractions to determine the purity and relative concentrations of the light chain proteins in each fraction. Desired fractions are those that contain concentrated light chain proteins without any visible protease bands. Combine the desired protein fractions and concentrate them using a 10,000 MWCO concentrating tube³⁵.
19. Flash freeze aliquots of protein in liquid nitrogen and immediately transfer to -80 °C for long-term storage (Figure 3B).
    NOTE: The yield of RLC and CALML4 using the E. coli His6-SUMO system is about 1-2 mg/L.

3. Myosin-7a-tailored actin filament gliding assay (3 h)

NOTE: The methods used in this section are similar to those described for other myosins³¹ with the major modifications being the incubation and application of myosin in high ionic strength buffer and the long interval required to achieve accurate measurement of frame-to-frame displacements.

1. Prepare 20 µM filamentous actin (F-actin) by polymerizing globular actin (G-actin) in polymerization buffer (50 mM KCl, 25 mM MOPS, 2 mM MgCl2, 1 mM DTT (pH 7.0)) at 4 °C overnight. Label F-actin by adding 1.2x molar excess rhodamine-phalloidin (see Table of Materials). Cover with aluminum foil and incubate on ice for at least 2 h.
   NOTE: G-actin can be purchased from commercial vendors (see Table of Materials) or purified from rabbit muscle acetone powder^31,37. Labeled F-actin can be stored at 4 °C for up to 2 months at this concentration.
2. Prepare a flow chamber as described previously³¹. Briefly, place two strips of double-sided tape 2-3 mm apart on a cleaned and nitrocellulose-treated microscope slide (see Table of Materials). Put the coverslip nitrocellulose side down onto the strips, creating a flow chamber with an approximate volume of 10 µL (Figure 4).
3. Performing actin filament gliding assay
   1. To the flow chamber, add one chamber volume of 0.2 mg/mL purified human myosin-7a protein in 500 mM NaCl Motility Buffer (500 mM NaCl, 20 mM MOPS, 5 mM MgCl₂, 0.1 mM EGTA (pH 7.4)). Let incubate for 5 min at room temperature.
      NOTE: It is critical to add myosin-7a in a high salt buffer as this relieves the autoinhibited state and allows myosin-7a to adhere to the surface in the activated conformation.
   2. Wash the flow chamber with three-chamber volumes of 1 mg/mL BSA (see Table of Materials) in 150 mM NaCl Motility Buffer (150 mM NaCl, 20 mM MOPS, 5 mM MgCl₂, 0.1 mM EGTA, 1 mM DTT (pH 7.4)) drawing the liquid through using a clean wipe at the outlet to absorb excess liquid. Incubate for 1 min after the 3^rd wash.
   3. Flow one chamber volume of 10 nM rhodamine phalloidin-labeled F-actin in 150 mM NaCl Motility Buffer through the flow chamber. Monitor binding of actin filaments to the surface under a fluorescent microscope with 100x objective. The density of actin filaments should be optimal (Figure 5A), ensuring enough filaments in the field-of-view for tracking yet sparse enough to prevent the overlapping of multiple filaments, which complicates tracking.
   4. Wash the flow chamber with three-chamber volumes of 150 mM NaCl Motility Buffer to remove unbound actin filaments and excess rhodamine-phalloidin.
   5. Initiate the motility by adding one chamber volume of the final buffer (200 mM NaCl, 20 mM MOPS, 5 mM MgCl₂, 0.1 mM EGTA, 50 mM DTT, 2 mM ATP, 2.5 mg/ml glucose, 100 µg/mL glucose oxidase, 2 µM RLC, 2 µM CaM, 2 µM CALML4).
   6. Record images on a fluorescence microscope using 561 nm excitation to visualize rhodamine phalloidin-labeled actin. Find an area on the slide with the desired actin density (Figure 5A) and capture images every 30 s for 30 min.
      NOTE: The velocity of mammalian myosin-7a is approximately 5 nm/s. When setting the acquisition frame rate, it is important to ensure that the displacements of the filaments between frames are long enough (more than one pixel) to allow for accurate tracking. Subpixel movements between frames can cause an overestimation of the speed. The frame rate of 30 s allows the actin filaments to move about 150 nm between frames, corresponding to more than two pixels on the microscope. However, care should be taken that significant displacement can be observed on the microscope being used. If available, multi-position recording can be used to simultaneously collect several field-of-view for analysis.
4. Analyzing the movement of the actin filaments using the FAST program (installed on a Mac)³⁸.
   NOTE: The FAST program used here is only one of many options for quantifying filament motion. We chose to use FAST because it is automated, fast, and does not require paid software. Other options include MATLAB-based algorithms such as FIESTA, ImageJ/FIJI-based methods such as MTrack2 and Manual Tracking, and Python-based methods such as Philament³⁹.
   1. Download and install the FAST program (github.com/NeilBillington/FASTrack3 – python3 compatible version with an additional module for. nd2 file conversion. NB: Original python2 version can be found at github.com/turalaksel/FASTrack).
   2. Run the FAST program as described in³⁸ using a tolerance value of 33 to retain smooth moving filaments only.
      NOTE: If images contain drift to mechanical issues with the stage or from collecting several series concurrently, the movies should first be stabilized. For this, we recommend the FIJI plugin Image Stabilizer, available for download at the following address – (https://www.cs.cmu.edu/~kangli/code/Image_Stabilizer.html).
   3. Use the FAST program to analyze the newly created .tif images by first setting the -xmax and -ymax values. These set the parameters for the scatter plot output by the program and represent the longest filament length (in nm) and the maximum filament velocity (in nm/s). We used 20,000 nm for -xmax and 20 nm/s for -ymax in this example to ensure all data is captured (Figure 5D).
   4. Use -px to set the pixel size of the acquired image, which here will be 65 nm, and -minv to set the minimum velocity (in nm/s) required of a filament for inclusion in the analysis. For the low velocity of myosin-7a in this example, use 0.1 nm/s.
   5. Use -maxd to set the maximum distance allowed between frames for filaments to be linked. This is set to avoid linking separate filaments between frames and in this example, we use the default 2000 nm.
   6. Use-pt to set the tolerance for fluctuations in filament velocities and only include filaments with smooth movements. This example uses a tolerance threshold of 33%. This means that filaments with a velocity standard deviation larger than 33% of the velocity mean will be excluded from further analysis.
   7. Lastly, use -d to indicate the folder containing the .tif stacks for analysis. The entire command string with the given parameters and values will be: fast -xmax 20000 -ymax 20 -px 65 -minv 0.1 -maxd 2000 -pt 33 -d [folder name containing movies].
      NOTE: The FAST program will output scatterplots (Figure 5D) with their associated raw data, which can be extracted for analysis and visualization in other programs (Figure 5C). It will also output a plot of the paths of the tracked filaments (Figure 5B), which should be used to verify that the tracking is working as expected. Examples of the plots and other possible data visualizations are included below.