We thank Dr. Fang Zhang for technical assistance with the preparation of the reagents used for collecting this data. This work was supported by the NHLBI/NIH Intramural Research Program funds HL001786 to J.R.S.

The authors declare no conflict of interest.

Presented here is a workflow for the purification and in vitro characterization of myosin 5a and nonmuscle myosin 2b. This set of experiments is useful for quantifying the mechanochemical properties of purified myosin constructs in a fast and reproducible manner. Although the two myosins shown here are just two specific examples out of the many possibilities, the conditions and techniques can be applied, with some tailoring, to most myosins and to many other motor proteins.
The protocols discussed here are subject to variations depending on the individual needs of the lab and experiments. For example, as discussed in the Expression and Molecular Biology section, the proteins used in this paper were generated from a co-infection of two or more viruses; however, successful protein expression can also be achieved with multi-expression vectors such as the p-FastBac dual expression vector or the BiGBac expression system53.
Several factors can hamper the successful production of the recombinant protein. To prevent protein degradation, it is imperative that every step of the protein purification is completed at the appropriate temperature, with all centrifugation steps occurring at 4 &#176;C and all other steps being performed on ice. Excess bands may be apparent in the gel of the dialyzed protein product. This could be indicative of degradation or contamination. Subsequent purification via size-exclusion or ionic-exchange chromatography can enhance the purity of the samples54. To that effect, it is recommended to save aliquots at each step of the protein purification process for troubleshooting, should there be a low yield of myosin or suspected protein contamination. Sometimes, there can be inadequate binding of the lysate to the resin in step 1.8, which can result in the loss of the myosin to the supernatant in subsequent centrifugations. This can be resolved by varying the duration of resin binding during this step, even leaving it to bind overnight, if necessary. Longer incubation times introduce a greater risk of protein degradation if contaminant proteases are not sufficiently inhibited, and myosins with proteolytically sensitive regions will be adversely affected. Additionally, improper washing of the resin both before and after use may result in the elution of an undesired protein product, so it is imperative that the proper protocols are followed before and after using the FLAG-affinity resin. If the resin is washed immediately after use and stored appropriately, it can be reused up to 20 times.
It is important to note that protein degradation also occurs during the expression stage and shortening expression times may be advantageous in terms of degradation, although this can be at the expense of the total yield. Following the procedures outlined here to monitor the protein sample at different stages of the protocol (i.e., before, during, and after purification) will help to determine the stages necessary for optimization. For myosins that repeatedly resist successful expression and purification, common problems can be co-expression with insufficient or inappropriate light chains as well as improper folding during overexpression. Appropriate light chains must be selected based on known interactions when possible and heavy chain to light chain baculovirus ratios must be tested in small-scale experiments to determine the optimum. For myosins that aggregate or yield little or no soluble active product, co-expression with chaperones can aid in successfully obtaining active protein54,55.
Purified myosin products inevitably contain a small population of damaged myosin, referred to as &#34;dead heads,&#34; which can be addressed in two ways. One method, outlined in this protocol, involves flowing unlabeled, or &#34;black&#34;, actin through the chamber in the gliding actin filament assay. Subsequently washing with ATP causes functional myosins to dissociate from the black actin while dead heads will remain bound to this unlabeled actin due to their inability to hydrolyze ATP and due to their high affinity. While performing the black actin wash, a syringe can be used to shear the actin effectively. Additional shearing can be accomplished by vortexing, provided that any resultant bubbles are removed by centrifugation. An alternative method is to selectively pellet the dead heads from the myosin sample by mixing myosin with F-actin and Mg-ATP at high salt (0.5 M) concentrations and sedimenting in a table-top ultracentrifuge. The myosins capable of hydrolyzing ATP under these conditions do not stay bound to the actin due to their low affinity for actin under these high salt conditions and are found in the supernatant, whereas the myosin dead heads remain bound to the actin in the pellet56. Similarly, a sedimentation with actin and resuspension of the pellet can also be used to remove myosins which are incapable of binding to actin in the absence of ATP. Note that a small proportion of this type of dead heads will have less of an impact on these types of assays. By doing a nucleotide-free sedimentation and resuspension followed by an ATP-bound centrifugation and resuspension, myosins that are competent for both actin-binding and ATP-dependent release from actin can be isolated.
Motility assays can also be modified in several ways. For example, in the gliding actin filament assay for the NM2b, the NM2b is phosphorylated in the chamber via the addition of MLCK, calmodulin, calcium, and ATP in the black actin step, as well as in the Final Buffer. However, the NM2b can also be phosphorylated in a tube, before performing the assay. By doing so, the percent of phosphorylated NM2b can be quantified by running a native gel with a urea-containing sample buffer or performing mass spectrometry57,58. The effect of temperature on myosin activity can also be investigated. This can be accomplished by employing an objective heating system on the microscope or an environmental enclosure, so that the flow cell is maintained at constant temperature. Ionic strength is another important consideration. For many myosins, actin affinity and enzymatic activity will be increased at lower ionic strength; for others, higher ionic strength is necessary59. In addition to providing valuable information about the myosin mechanism, lowering the ionic strength can enhance motility and make myosin more accessible to investigation with many assays. In contrast, some motors will exhibit electrostatic tethering effects, which will slow motility at lower ionic strengths. Finally, when assaying the movement of NM2b filaments, it is crucial to maintain ionic strength within a narrow range (150-200 mM ionic strength), approximating those found in most cell types. The use of lower ionic strengths results in aggregation of the myosin filaments, while the filaments depolymerize at higher ionic strengths.
With many myosins, particularly those with low duty ratios, the conditions of the final buffer given for M5a would result in the fluorescently labeled actin filaments being only loosely bound to the surface or dissociating altogether. This results in erratic movements that complicate quantification. Better quality movement can often be obtained using methylcellulose (0.7%) in the Final Buffer. Methylcellulose is a viscous crowding agent and forces actin filaments to remain close to the surface even when the density of attached myosin motors is sparse60. Similarly, it has been observed that the inclusion of methylcellulose in the final buffer of the single filament motility assay is necessary to observe movement with NM2a, and the same phenomenon was reported for smooth muscle myosin filaments29,61. This also increases the processivity of NM2b filaments. One potentially unwanted side effect of using methylcellulose in this assay is that the crowding agent properties can promote the lateral association of myosin filaments into bundles. Alternatives to methylcellulose when troubleshooting a lack of movement or loosely bound actin filaments in the gliding actin filament assay are to lower the salt concentration in the motility buffers or to increase the myosin surface density. As stated above, a high ionic strength in the motility buffers has been shown to lower the ability of some myosins to bind to actin29,34,62.
Another variation of the gliding actin filament assay is the use of antibodies to anchor the myosin onto the glass coverslip. For example, if a GFP is present at the C-terminal end of the myosin construct, an anti-GFP antibody can be used to fix the GFP-myosin to the coverslip36,63,64. This can aid with obtaining successful motility in situations where the geometry of the system may otherwise hamper actin translocation, such as in the case of testing artificial or short lever arms54,64. Additionally, the effect of load on translocation velocities can be investigated in the gliding actin filament assay by employing actin-binding proteins such as &#945;-actinin or utrophin39,50,65. Such a measurement can be useful to compare the effect of load on an ensemble of myosins versus the load-dependent kinetics of a single myosin that can be measured using an optical trapping assay66,67. This can be accomplished by adding increasing amounts of an actin-binding protein along with the myosin in the initial step. The actin-binding protein binds to the surface and exerts a frictional load on the actin filaments that are being moved by myosins, which results in a graded velocity as the concentration of actin-binding protein on the surface is increased39.
The single molecule/ensemble motility assay can also be adapted to investigate the effect of various actin structures on myosin movement. For example, rather than observing myosin movement on top of single actin filaments, fascin- or &#945;-actinin-mediated actin bundles can be studied as an in vitro reconstitution of the actin filament network found in cells68,69. The effect of actin-binding proteins such as tropomyosin can also be studied65,70,71,72.
Of note is the versatility in choosing a label for the single molecule/ensemble motility assay. In this report, a GFP label was used on both the M5a-HMM and NM2b; however, many other labels can be used. Examples include HaloTag or SNAP-tag, which can be genetically fused to the myosin and covalently bind a synthetic dye. The benefit of HaloTag technology lies in its versatility for several experimental adaptations, such as labeling with different colors or adding a biotin affinity tag29,73. Additionally, the use of quantum dot technology can be employed to improve the resolution of single molecule fluorescence tracking, which also addresses the limitation of GFP&#39;s low brightness and the tendency for photobleaching74. Tags can be successfully attached to light chains as well as the heavy chain11,75,76.
For achieving success in the single molecule TIRF motility assay, a key factor is using a well blocked and functionalized surface. A simple method to achieve moderate blocking is to use biotinylated-BSA bound to a nitrocellulose surface. Although this will work well enough to characterize many motors, including M5a, the level of nonspecific binding on such a surface is prohibitive for reproducing clean movement with samples such as NM2b. A key breakthrough in this regard was the transition to PEGylated surfaces doped with biotin-PEG for functionalization77. The PEG surfaces provide a far superior level of surface blocking and a defect-free PEGylated surface can remain free of nonspecific binding for very long periods of time. The specific protocol detailed here allows the production of biotinylated PEG surfaces in a matter of hours and if immediately stored as described, the surfaces can be used for several weeks with only a marginal decline in quality.
A key consideration before collecting data for tracking is the acquisition frame rate. Movement between subsequent frames must be large enough to avoid oversampling errors. High sampling rates will yield overestimated velocities due to the division of localization errors by a small time interval and increase the apparent error of the measurement. In cases where the raw data is too finely sampled, the data can be down-sampled by taking every Nth frame to create a new stack and considering the change in frame rate that results. Subpixel movements between frames must be avoided and movements of several hundred nanometers are required to obtain accurate values. In all cases where a new sample is being characterized, the results generated by automated analysis must be compared to a small dataset of manually tracked filaments for consistency.
When analyzing data from single molecule motility experiments, care must be taken when choosing which parameters to measure, how to filter data, and how to fit data. As stated above, the sampling rate can be an important factor when analyzing velocity data. For many myosins, processive runs will be short and well approximated by a straight line. In such cases, the start to endpoint distance of the track may provide a good measure of the run length and this can be divided by the duration of the track to yield a good estimate of velocity. In cases where the tracks are very long and follow curved paths around bent filaments, this type of analysis will yield inaccurate results and a total distance traveled must be used, using an acquisition rate that allows for successive localization points to be sufficiently well spaced to avoid oversampling errors as described above, while being close enough together that the straight line distance between them remains a good approximation of the curve between those points. In addition, for motor proteins with long run lengths in relation to the length of the track, additional statistics such as the Kaplan-Meier estimator must be made when calculating run lengths78. The same is true for situations in which photobleaching is sufficiently likely to occur before the end of a processive run. Another phenomenon that can be observed in single molecule fluorescence studies is photoblinking, in which fluorophores switch between the on and off state rapidly and appear to blink. This typically does not occur in these motility experiments; however, if this does occur, the laser intensity and exposure times can be decreased which should minimize the effect. Several chemicals, including &#946;-mercaptoethanol, Trolox, cyclooctateraene, n-propyl gallate, 4-nitrobenzyl alcohol, and 1,4-diazabicyclo[2.2.2]octane can be utilized to mitigate this as well79.
In summary, this article presents detailed protocols that are robust in their ability to quantify mechanochemical properties such as actin translocation velocity, myosin translocation velocity, and myosin run length. These assays are reproducible and can be used to determine the quality of the purified myosin even in situations where the motile characteristics are not the specific end goal of the study. In addition, changes such as pH, temperature, and chemical regulators can be introduced to these assays to examine how the mechanochemistry of the studied myosin is affected. Taken together, the actin gliding and inverted motility assays can allow for a better understanding of myosin ensemble behavior and intermolecular variations in molecular motor mechanics and kinetics. The fluorescence microscopy-based assays described here support a reductionist&#39;s approach to cytoskeletal research and can be a powerful tool to understand protein-protein dynamics in vitro. Together, data collected from these highly controlled experiments can be used to advise mechanobiologists of key actomyosin behaviors that may be relevant at the cell biological level, and beyond.

Myosins are motor proteins that exert force on actin filaments using the energy derived from adenosine triphosphate (ATP) hydrolysis1. Myosins contain a head, neck, and tail domain. The head domain contains the actin-binding region as well as the site of ATP binding and hydrolysis. The neck domains are composed of IQ motifs, which bind to light chains, calmodulin, or calmodulin-like proteins2,3. The tail region has several functions specific to each class of myosins, including but not limited to the dimerization of two heavy chains, binding of cargo molecules, and regulation of the myosin via autoinhibitory interactions with the head domains1.
The motile properties of myosin vary greatly between classes. Some of these properties include duty ratio (the fraction of myosin&#39;s mechanical cycle in which the myosin is bound to actin) and processivity (the ability of a motor to make multiple steps on its track before detachment)4. The over 40 classes of myosins were determined based on sequence analyses5,6,7,8. The class 2 myosins are classified as &#34;conventional&#34; since they were the first to be studied; all other classes of myosins are, therefore, classified as &#34;unconventional.&#34;
Myosin 5a (M5a) is a class 5 myosin and is a processive motor, meaning that it can take multiple steps along actin before dissociating. It has a high duty ratio, indicating that it spends a large part of its mechanical cycle bound to actin9,10,11,12,13,14. In common with other myosins, the heavy chain contains an N-terminal motor domain that includes both an actin-binding and an ATP hydrolysis site followed by a neck region that serves as a lever-arm, with six IQ motifs that bind to essential light chains (ELC) and calmodulin (CaM)15. The tail region contains &#945;-helical coiled-coils, which dimerize the molecule, followed by a globular tail region for binding cargo. Its kinetics reflect its involvement in the transport of melanosomes in melanocytes and of the endoplasmic reticulum in Purkinje neurons16,17. M5a is considered the prototypical cargo transport motor18.
Class 2 myosins, or the conventional myosins, include the myosins that power contraction of skeletal, cardiac, and smooth muscle in addition to the nonmuscle myosin 2 (NM2) isoforms, NM2a, 2b, and 2c19. The NM2 isoforms are found in the cytoplasm of all cells and have shared roles in cytokinesis, adhesion, tissue morphogenesis, and cell migration19,20,21,22. This paper discusses conventional myosin protocols in the context of nonmuscle myosin 2b (NM2b)23. NM2b, in comparison to M5a, has a low duty ratio and is enzymatically slower with a Vmax of 0.2 s-1 23 compared to M5a&#39;s Vmax of &#8776;18 s-1 24. Notably, truncated NM2b constructs with two heads do not readily move processively on actin; rather, each encounter with actin results in a power stroke followed by dissociation of the molecule25.
NM2b contains two myosin heavy chains, each with one globular head domain, one lever-arm (with one ELC and one regulatory light chain (RLC)), and an &#945;-helical coiled-coil rod/tail domain, approximately 1,100 amino acids long, that dimerizes these two heavy chains. The enzymatic activity and structural state of NM2b are regulated by phosphorylation of the RLC23. Unphosphorylated NM2b, in the presence of ATP and physiological ionic strengths (approximately 150 mM salt), adopts a compact conformation wherein the two heads make participate in an asymmetric interaction and the tail folds back over the heads in two places23. In this state, the myosin does not interact strongly with actin and has very low enzymatic activity. Upon RLC phosphorylation by calmodulin-dependent myosin light chain kinase (MLCK) or Rho-associated protein kinase, the molecule extends and associates with other myosins through the tail region to form bipolar filaments of approximately 30 myosin molecules23. The aforementioned phosphorylation of the RLC also leads to increased actin-activated ATPase activity of NM2b by approximately four times26,27,28. This bipolar filament arrangement, featuring many myosin motors at each end, is optimized for roles in contraction and tension maintenance, where actin filaments with opposing polarities can be moved relative to each other23,29. Accordingly, NM2b has been shown to act as an ensemble of motors when interacting with actin. The large number of motors within this filament allow NM2b filaments to move processively on actin filaments, making in vitro&#160;filament processivity possible to characterize29.
While progress has been made in understanding the role of myosins in the cell, there is a need to understand their individual characteristics at the protein level. To understand actomyosin interactions at a simple protein-protein interaction level, rather than inside of a cell, we can express and purify recombinant myosins for use in in vitro studies. The results of such studies then inform mechanobiologists about the biophysical properties of specific myosins that ultimately drive complex cellular processes12,13,14,25,29. Typically, this is accomplished by adding an affinity tag to a full-length or truncated myosin construct and purifying via affinity chromatography29,30,31. Additionally, the construct can be engineered to include a genetically encodable fluorophore or a tag for protein labeling with a synthetic fluorophore. By adding such a fluorescent label, single molecule imaging studies can be performed to observe myosin mechanics and kinetics.
Following purification, the myosin can be characterized in several ways. ATPase activity can be measured by colorimetric methods, providing insight into the overall energy consumption and actin affinity of the motor under different conditions32. To learn about the mechanochemistry of its motility, further experiments are required. This paper details two in vitro fluorescence microscopy-based methods that can be used to characterize the motile properties of a purified myosin protein.
The first of these methods is the gliding actin filament assay, which can be used to quantitatively study the ensemble properties of myosin motors, as well as qualitatively study the quality of a batch of purified protein33. Although this paper discusses the use of total internal reflection fluorescence (TIRF) microscopy for this assay, these experiments can be effectively performed using a wide-field fluorescence microscope equipped with a digital camera, commonly found in many labs34. In this assay, a saturating layer of myosin motors is attached to a coverslip. This can be accomplished using nitrocellulose, antibodies, membranes, SiO2-derivatized surfaces (such as trimethylchlorosilane), among others29,33,35,36,37,38. Fluorescently labeled actin filaments are passed through the coverslip chamber, upon which the actin binds to the myosin attached to the surface. Upon addition of ATP (and kinases in the study of NM2), the chamber is imaged to observe the translocation of actin filaments by the surface-bound myosins. Tracking software can be used to correlate the velocity and length of each gliding actin filament. Analysis software can also provide a measure of the number of both moving and stationary actin filaments, which can be useful to determine the quality of a given myosin preparation. The proportion of stalled filaments can also be intentionally modulated by surface tethering of actin to other proteins and measured to determine the load dependence of the myosin39. Because each actin filament can be propelled by a large number of available motors, this assay is very reproducible, with the final measured velocity being robust to perturbations such as alterations in the starting myosin concentration or the presence of additional factors in the solution. This means it can be easily modified to study myosin activity under different conditions, such as altered phosphorylation, temperature, ionic strength, solution viscosity, and the effects of load induced by surface tethers. Although factors such as strong-binding myosin &#34;dead heads&#34; incapable of ATP hydrolysis can cause stalled actin filaments, multiple methods exist to mitigate such issues and allow for accurate measurements. The kinetic properties of myosin vary greatly across classes and, depending on the specific myosin used, the speed of actin filament gliding in this assay can vary from under 20 nm/s (myosin 9)40,41, and up to 60,000 nm/s (Characean myosin 11)42.
The second assay inverts the geometry of the gliding actin filament assay12. Here, the actin filaments are attached to the coverslip surface and the movement of single molecules of M5a or of individual bipolar filaments of NM2b are visualized. This assay can be used to quantify the run lengths and velocities of single myosin molecules or filaments on actin. A coverslip is coated with a chemical compound that blocks non-specific binding and simultaneously functionalizes the surface, such as biotin-polyethylene glycol (biotin-PEG). The addition of modified avidin derivatives then primes the surface and biotinylated actin is passed through the chamber, resulting in a layer of F-actin stably bound to the bottom of the chamber. Finally, activated and fluorescently labeled myosin (typically 1-100 nM) is flowed through the chamber, which is then imaged to observe myosin movement over the stationary actin filaments.
These modalities represent fast and reproducible methods that can be employed to examine the dynamics of both nonmuscle and muscle myosins. This report outlines the procedures to purify and characterize both M5a and NM2b, representing unconventional and conventional myosins, respectively. This is followed by a discussion of some of the myosin-specific adaptations, which can be performed to achieve successful capturing of motion in the two types of the assay.
Expression and molecular biology 
The cDNA for the myosin of interest must be cloned onto a modified pFastBac1 vector that encodes for either a C-terminal FLAG-tag (DYKDDDDK) if expressing M5a-HMM, or an N-terminal FLAG-tag if expressing the full-length molecule of NM2b23,43,44,45,46. C-terminal FLAG-tags on NM2b results in a weakened affinity of the protein for the FLAG-affinity column. In contrast, the N-terminally FLAG-tagged protein usually binds well to the FLAG-affinity column23. The N-terminally tagged protein retains enzymatic activity, mechanical activity and phosphorylation-dependent regulation23.
In this paper, a truncated mouse M5a heavy meromyosin (HMM)-like construct with a GFP between the FLAG-tag and the C-terminus of the myosin heavy chain was used. Note that unlike NM2b, M5a-HMM can be successfully tagged and purified with either N- or C-terminal FLAG tags and in both cases the resulting construct will be active. The M5a heavy chain was truncated at amino acid 1090 and contains a three amino acid linker (GCG) between the GFP and the coiled-coil region of the M5a47. No additional linker was added between the GFP and FLAG-tag. M5a-HMM was co-expressed with calmodulin. The full-length human NM2b construct was co-expressed with ELC and RLC. The N-termini of the RLC was fused with a GFP via a linker of five amino acids (SGLRS). Directly attached to the FLAG-tag was a HaloTag. Between the HaloTag and the N-terminus of the myosin heavy chain was a linker made of two amino acids (AS).
Both myosin preparations were purified from one liter of Sf9 cell culture infected with baculovirus at a density of approximately 2 x 106 cells/mL. The volumes of the baculovirus for each subunit depended on the virus&#39;s multiplicity of infection as determined by the manufacturer&#39;s instructions. In the case of M5a, cells were co-infected with two different baculoviruses-one for calmodulin, and one for M5a heavy chain. In the case of the NM2b, cells were co-infected with three different viruses-one for ELC, one for RLC, and one for NM2b heavy chain. For labs working with a diversity of myosins (or other multi-complex proteins), this approach is efficient since it allows for many combinations of heavy and light chains and commonly used light chains such as calmodulin can be co-transfected with many different myosin heavy chains. All cell work was completed in a biosafety cabinet with proper sterile technique to avoid contamination.
For the expression of both M5a and NM2b, the Sf9 cells producing the recombinant myosins were collected 2-3 days post-infection, via centrifugation, and stored at -80 &#176;C. Cell pellets were obtained by centrifuging the co-infected Sf9 cells at 4 &#176;C for 30 min at 2,800 x g. The protein purification process is detailed below.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 mL Syringe</td>
			<td>BD</td>
			<td>309628</td>
			<td></td>
		</tr>
		<tr>
			<td>2 M CaCl2 Solution</td>
			<td>VWR</td>
			<td>10128-558</td>
			<td></td>
		</tr>
		<tr>
			<td>2 M MgCl2 Solution</td>
			<td>VWR</td>
			<td>10128-298</td>
			<td></td>
		</tr>
		<tr>
			<td>27 Gauge Needle</td>
			<td>Becton Dickinson</td>
			<td>309623</td>
			<td></td>
		</tr>
		<tr>
			<td>5 M NaCl Solution</td>
			<td>KD Medical</td>
			<td>RGE-3270</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetic Acid</td>
			<td>ThermoFisher Scientific</td>
			<td>984303</td>
			<td></td>
		</tr>
		<tr>
			<td>Amyl Acetate</td>
			<td>Ladd Research Industries</td>
			<td>10825</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-FLAG M2 Affinity Gel</td>
			<td>Millipore Sigma</td>
			<td>A2220</td>
			<td>https://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/Bulletin/a2220bul.pdf</td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>Millipore Sigma</td>
			<td>A7699</td>
			<td></td>
		</tr>
		<tr>
			<td>Biotinylated G-Actin</td>
			<td>Cytoskeleton, Inc.</td>
			<td>AB07</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Millipore Sigma</td>
			<td>5470</td>
			<td></td>
		</tr>
		<tr>
			<td>bPEG-silane</td>
			<td>Laysan Bio, Inc</td>
			<td>Biotin-PEG-SIL-3400-1g</td>
			<td></td>
		</tr>
		<tr>
			<td>Bradford Reagent Concentrate</td>
			<td>Bio-Rad</td>
			<td>5000006</td>
			<td></td>
		</tr>
		<tr>
			<td>Calmodulin</td>
			<td></td>
			<td>PMID: 2985564</td>
			<td></td>
		</tr>
		<tr>
			<td>Catalase</td>
			<td>Millipore Sigma</td>
			<td>C40</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Line (Sf9) in SF-900 II SFM</td>
			<td>ThermoFisher Scientific</td>
			<td>11496015</td>
			<td>http://tools.thermofisher.com/content/sfs/manuals/bevtest.pdf https://tools.thermofisher.com/content/sfs/manuals/bactobac_man.pdf</td>
		</tr>
		<tr>
			<td>Circular Filter Paper - Gliding Assay</td>
			<td>Millipore Sigma</td>
			<td>WHA1001125</td>
			<td></td>
		</tr>
		<tr>
			<td>Circular Filter Paper - Inverted Assay</td>
			<td>Millipore Sigma</td>
			<td>WHA1001090</td>
			<td></td>
		</tr>
		<tr>
			<td>cOmplete, EDTA-Free Protease Inhibitor Tablets</td>
			<td>Millipore Sigma</td>
			<td>5056489001</td>
			<td>This should be stored at 4 &#176;C. The tablets can be used directly or can be reconstituted as a 25x stock solution by dissolving 1 tablet in 2 mL of distilled water. The resulting solution can be stored at 4 &#176;C for 1-2 weeks or at least 12 weeks at -20 &#176;C.&#160;</td>
		</tr>
		<tr>
			<td>Concentrating Tubes (100,000 MWCO)</td>
			<td>EMD Millipore Corporation</td>
			<td>UFC910024</td>
			<td>The MWCO of the tube is not necessarily &#34;one size fits all,&#34; as long as the MWCO is less than the total molecular weight of the protein being purified. The NM2b herein was concentrated with a 100,000 MWCO tube and the M5a was concentrated with a 30,000 MWCO tube.</td>
		</tr>
		<tr>
			<td>Coomassie Brilliant Blue R-250 Dye</td>
			<td>ThermoFisher Scientific</td>
			<td>20278</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslip Rack</td>
			<td>Millipore Sigma</td>
			<td>Z688568-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips: Gliding Acting Filament Assay</td>
			<td>VWR International</td>
			<td>48366-227</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips: Inverted Motility Assay</td>
			<td>Azer Scientific</td>
			<td>ES0107052</td>
			<td></td>
		</tr>
		<tr>
			<td>Dialysis Tubing (3500 Dalton MCWO)</td>
			<td>Fischer Scientific</td>
			<td>08-670-5A</td>
			<td>The diameter of the dialysis tube can vary, but the MWCO should be the same. The NM2b used herein was dialyzed in an 18 mm dialysis tube. The tubes can be stored in 20% alcohol solution at 4 &#176;C.</td>
		</tr>
		<tr>
			<td>DL-Dithiothreitol</td>
			<td>Millipore Sigma</td>
			<td>D0632</td>
			<td></td>
		</tr>
		<tr>
			<td>Double-Sided Tape</td>
			<td>Office Depot</td>
			<td>909955</td>
			<td></td>
		</tr>
		<tr>
			<td>DYKDDDDK Peptide</td>
			<td>GenScript</td>
			<td>RP10586</td>
			<td>This can be dissolved in a buffer containing 0.1 M NaCl, 0.1 mM EGTA, 3 mM NaN3, and 10 mM MOPS (pH 7.2) to a final concentration of 50 mg/mL. This can be stored at -20 &#176;C as 300 &#181;L aliquots.&#160;</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Millipore Sigma</td>
			<td>E4378</td>
			<td></td>
		</tr>
		<tr>
			<td>Elution Column</td>
			<td>Bio-Rad</td>
			<td>761-1550</td>
			<td>These can be reused. To clean, rinse the column with 2-3 column volumes of PBS and distilled water. Chill the column at 4&#176; C before use.</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Fischer Scientific</td>
			<td>A4094</td>
			<td></td>
		</tr>
		<tr>
			<td>G-actin</td>
			<td></td>
			<td>PMID: 4254541</td>
			<td>G-actin stock can be stored at 200 &#956;M in liquid N2.</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Millipore Sigma</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose Oxidase</td>
			<td>Millipore Sigma</td>
			<td>G2133</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine Buffer Solution, 100 mM, pH 2-2.5, 1 L</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-295018</td>
			<td></td>
		</tr>
		<tr>
			<td>HaloTag</td>
			<td>Promega</td>
			<td>G100A</td>
			<td></td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>Millipore Sigma</td>
			<td>320331</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fischer Scientific</td>
			<td>P217-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Large-Orifice Pipet Tips</td>
			<td>Fischer Scientific</td>
			<td>02-707-134</td>
			<td></td>
		</tr>
		<tr>
			<td>Leupeptin Protease Inhibitor</td>
			<td>ThermoFisher Scientific</td>
			<td>78435</td>
			<td></td>
		</tr>
		<tr>
			<td>Mark12 Unstained Standard Ladder</td>
			<td>ThermoFisher Scientific</td>
			<td>LC5677</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Millipore Sigma</td>
			<td>MX0482</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylcellulose</td>
			<td>Millipore Sigma</td>
			<td>M0512</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fischer Scientific</td>
			<td>12-553-10</td>
			<td></td>
		</tr>
		<tr>
			<td>MOPS</td>
			<td>Fischer Scientific</td>
			<td>BP308-100</td>
			<td></td>
		</tr>
		<tr>
			<td>mPEG-silane</td>
			<td>Laysan Bio, Inc</td>
			<td>MPEG-SIL-2000-1g</td>
			<td></td>
		</tr>
		<tr>
			<td>Myosin Light Chain Kinase</td>
			<td></td>
			<td>PMID: 23148220</td>
			<td>FLAG-tagged MLCK can be purified the same way that the FLAG-tagged myosin was purified herein.&#160;</td>
		</tr>
		<tr>
			<td>NaN3</td>
			<td>Millipore Sigma</td>
			<td>S8032</td>
			<td></td>
		</tr>
		<tr>
			<td>NeutrAvidin</td>
			<td>ThermoFisher Scientific</td>
			<td>31050</td>
			<td></td>
		</tr>
		<tr>
			<td>Nitrocellulose</td>
			<td>Ladd Research Industries</td>
			<td>10800</td>
			<td></td>
		</tr>
		<tr>
			<td>NuPAGE 4 to 12% Bis-Tris Mini Protein Gel, 15-well</td>
			<td>ThermoFisher Scientific</td>
			<td>NP0323PK2</td>
			<td></td>
		</tr>
		<tr>
			<td>NuPAGE LDS Sample Buffer (4X)</td>
			<td>ThermoFisher Scientific</td>
			<td>NP0007</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-Buffered Saline, pH 7.4</td>
			<td>ThermoFisher Scientific</td>
			<td>10010023</td>
			<td></td>
		</tr>
		<tr>
			<td>PMSF</td>
			<td>Millipore Sigma</td>
			<td>78830</td>
			<td>PMSF can be made as a 0.1 M stock solution in isopropanol and stored in 4 &#176;C. Isopropanol addition results in crystal precipitation, which can be dissolved by stirring at room temperature. Immediately before use, PMSF can be added dropwise to a rapidly stirring solution to a final concentration of 0.1 mM.&#160;</td>
		</tr>
		<tr>
			<td>Razor Blades</td>
			<td>Office Depot</td>
			<td>397492</td>
			<td></td>
		</tr>
		<tr>
			<td>Rhodamine-Phalloidin</td>
			<td>ThermoFisher Scientific</td>
			<td>R415</td>
			<td>Stock can be diluted in 100% methanol to a final concentration of 200 &#956;M.</td>
		</tr>
		<tr>
			<td>Sf9 Media</td>
			<td>ThermoFisher Scientific</td>
			<td>12658-027</td>
			<td>This should be stored at 4&#176; C. Its shelf life is 18 months from the date of manufacture.</td>
		</tr>
		<tr>
			<td>Tissue Culture Dish - Gliding Assay</td>
			<td>Corning</td>
			<td>353025</td>
			<td>Each tissue culture dish can hold approximately nine coverslips.</td>
		</tr>
		<tr>
			<td>Tissue Culture Dish - Inverted Assay</td>
			<td>Corning</td>
			<td>353003</td>
			<td>Each tissue culture dish can hold approximately four coverslips.</td>
		</tr>
		<tr>
			<td>Smooth-sided 200 &#181;L Pipette Tips</td>
			<td>Thomas Scientific</td>
			<td>1158U38</td>
			<td></td>
		</tr>
		<tr>
			<td>EQUIPMENT</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>ThermoFisher Scientific</td>
			<td>75006590</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon</td>
			<td>Model: Eclipse Ti with H-TIRF system with 100x TIRF Objective (N.A. 1.49)</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Camera</td>
			<td>Andor</td>
			<td>Model: iXon DU888 EMCCD camera (1024 x 1024 sensor format)</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Environmental Control Box</td>
			<td>Tokai HIT</td>
			<td>Custom Thermobox</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Laser Unit</td>
			<td>Nikon</td>
			<td>LU-n4 four laser unit with solid state lasers for 405nm, 488nm, 561nm,and 640nm</td>
			<td></td>
		</tr>
		<tr>
			<td>Mid Bench Centrifuge</td>
			<td>ThermoFisher Scientific</td>
			<td>Model: CR3i</td>
			<td></td>
		</tr>
		<tr>
			<td>Misonix Sonicator</td>
			<td>Misonix</td>
			<td>XL2020</td>
			<td></td>
		</tr>
		<tr>
			<td>Optima Max-Xp Tabletop Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td>393315</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasma-Cleaner</td>
			<td>Diener electronic GmbH + Co. KG</td>
			<td>System Type: Zepto</td>
			<td></td>
		</tr>
		<tr>
			<td>Sonicator Probe (3.2 mm)</td>
			<td>Qsonica</td>
			<td>4418</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard Incubator</td>
			<td>Binder</td>
			<td>Model: 56</td>
			<td></td>
		</tr>
		<tr>
			<td>Waverly Tube Mixer</td>
			<td>Waverly</td>
			<td>TR6E</td>
			<td></td>
		</tr>
		<tr>
			<td>SOFTWARE</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ FIJI</td>
			<td>https://imagej.net/Fiji/Downloads</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>FAST (Version 1.01)</td>
			<td>http://spudlab.stanford.edu/fast-for-automatic-motility-measurements</td>
			<td>FAST is available for Mac OSX and Linux based systems.</td>
			<td></td>
		</tr>
		<tr>
			<td>Image Stabilizer Plugin</td>
			<td>https://imagej.net/Image_Stabilizer</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ TrackMate</td>
			<td>https://imagej.net/TrackMate</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging Software</td>
			<td>NIS Elements (AR package)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>http://www.cs.cmu.edu/~kangli/code/Image_Stabilizer.html</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>File:TrackMate-manual.pdf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>https://github.com/turalaksel/FASTrack</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>https://github.com/turalaksel/FASTrack/blob/master/README.md</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

myosin-specific adaptations of in vitro fluorescence microscopy-based motility assays

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic properties are adapted for the particular function they execute in cells. Myosin 5a processively walks on F-actin to transport melanosomes and vesicles in cells. Conversely, nonmuscle myosin 2b operates as a bipolar filament containing approximately 30 molecules. It moves F-actin of opposite polarity toward the center of the filament, where the myosin molecules work asynchronously to bind actin, impart a power stroke, and dissociate before repeating the cycle. Nonmuscle myosin 2b, along with its other nonmuscle myosin 2 isoforms, has roles that include&#160;cell adhesion, cytokinesis, and tension maintenance. The mechanochemistry of myosins can be studied by performing in vitro motility assays using purified proteins. In the gliding actin filament assay, the myosins are bound to a microscope coverslip surface and translocate fluorescently labeled F-actin, which can be tracked. In the single molecule/ensemble motility assay, however, F-actin is bound to a coverslip and the movement of fluorescently labeled myosin molecules on the F-actin is observed. In this report, the purification of recombinant myosin 5a from Sf9 cells using affinity chromatography is outlined. Following this, we outline two fluorescence microscopy-based assays: the gliding actin filament assay and the inverted motility assay. From these assays, parameters such as actin translocation velocities and single molecule run lengths and velocities can be extracted using the image analysis software. These techniques can also be applied to study the movement of single filaments of the nonmuscle myosin 2 isoforms, discussed herein in the context of nonmuscle myosin 2b. This workflow represents a protocol and a set of quantitative tools that can be used to study the single molecule and ensemble dynamics of nonmuscle myosins.

Watch this Scientific Journal Video about Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays at JoVE.com

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Myosin proteins bind and interact with filamentous actin (F-actin) and are found in organisms across the phylogenetic tree. Their structure and enzymatic ...

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Unit 1

Cell Polarization and Migration

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5.7K Views.  National Institutes of Health. Presented here is a procedure to express and purify myosin 5a followed by a discussion of its characterization, using both ensemble and single molecule in vitro fluorescence microscopy-based assays, and how these methods can be modified for the characterization of nonmuscle myosin 2b.

Video: Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

Automated Acoustic Dispensing for the Serial Dilution of Peptide Agonists in Potency Determination Assays

As with small molecule drug discovery, screening for peptide agonists requires the serial dilution of peptides to produce concentration-response curves. Screening peptides affords an additional layer of complexity as conventional tip-based sample handling methods expose peptides to a large surface area of plasticware, providing an increased opportunity for peptide loss via adsorption. Preventing excessive exposure to plasticware reduces peptide loss via adherence to plastics and thus minimizes inaccuracies in potency prediction, and we have previously described the benefits of non-contact acoustic dispensing for in vitro high-throughput screening of peptide agonists1. Here we discuss a fully integrated automation solution for non-contact acoustic preparation of peptide serial dilutions in microtiter plates utilizing the example of screening for peptide agonists at the mouse glucagon-like peptide-1 receptor (GLP-1R). Our methods allow for high-throughput cell-based assays to screen for agonists and are easily scalable to support increased sample throughput, or to allow for increased numbers of assay plate copies (e.g., for a panel of more target cell lines).

Peptide adsorption to plasticware during traditional tip-based serial dilutions can significantly impact potency determination and confound the understanding of structure-activity relationships used for lead identification and lead optimization phases of drug discovery. Here methods for automated acoustic non-contact serial dilution of peptide samples are described.

As with small molecule drug discovery, screening for peptide agonists requires the serial dilution of peptides to produce concentration-response curves. ...

In Vitro Polymerization of F-actin on Early Endosomes

Many early endosome functions, particularly cargo protein sorting and membrane deformation, depend on patches of short F-actin filaments nucleated onto the endosomal membrane. We have established a microscopy-based in vitro assay that reconstitutes the nucleation and polymerization of F-actin on early endosomal membranes in test tubes, thus rendering this complex series of reactions amenable to genetic and biochemical manipulations. Endosomal fractions are prepared by floatation in sucrose gradients from cells expressing the early endosomal protein GFP-RAB5. Cytosolic fractions are prepared from separate batches of cells. Both endosomal and cytosolic fractions can be stored frozen in liquid nitrogen, if needed. In the assay, the endosomal and cytosolic fractions are mixed, and the mixture is incubated at 37 &#176;C under appropriate conditions (e.g., ionic strength, reducing environment). At the desired time, the reaction mixture is fixed, and the F-actin is revealed with phalloidin. Actin nucleation and polymerization are then analyzed by fluorescence microscopy. Here, we report that this assay can be used to investigate the role of factors that are involved either in actin nucleation on the membrane, or in the subsequent elongation, branching, or crosslinking of F-actin filaments.

In Vitro Polymerization of F-actin on Early Endosomes

Early endosome functions depend on F-actin polymerization. Here, we describe a microscopy-based in vitro assay that reconstitutes the nucleation and polymerization of F-actin on early endosomal membranes in test tubes, thus rendering this complex series of reactions amenable to biochemical and genetic manipulations.

Many early endosome functions, particularly cargo protein sorting and membrane deformation, depend on patches of short F-actin filaments nucleated onto ...

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine whether there is an imbalance in age-related energy metabolism between mitochondrial oxidative phosphorylation and aerobic glycolysis. Here, we show the utilization of commercial colorimetric assay kits for lactate and pyruvate in the model organism C. elegans. Recently, the sensitivity and accuracy of the colorimetric/fluorimetric assay kits have been improved greatly by the research and development conducted by reagent manufacturers. The improved reagents have enabled the use of small-scale assays with a 96-well plate in C. elegans. In general, a fluorimetric assay is superior in sensitivity to a colorimetric assay; however, the colorimetric approach is more suitable for the use in common laboratories. Another important issue in these assays for quantitative determination is protein precipitation of homogenized C. elegans samples. In our protein precipitation method, common precipitants (e.g., trichloroacetic acid, perchloric acid and metaphosphoric acid) are used for sample preparation. A protein-free assay sample is prepared by directly adding cold precipitant (final concentration of 5%) during homogenization.

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

We describe a modified small-scale extraction and colorimetric assays of lactate and pyruvate in the nematode C. elegans. When utilizing commercial assay kits, the technical development of their sensitivity and accuracy is important. Protein precipitation in extraction is the most critical step for the quantitative determination of intracellular metabolites.

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine ...

Quantification of Hypopigmentation Activity In Vitro

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is synthesized in response to multiple cellular and environmental factors. Melanin protects skin cells from ultraviolet damage, but also has biophysical and biochemical functions. Excessive production or accumulation of melanin in melanocytes can cause dermatological problems, such as freckles, dark spots, melasma, and moles. Therefore, the control of melanogenesis with hypopigmentation agents is important in individuals with clinical or cosmetic needs. Melanin is primarily synthesized in the melanosomes of melanocytes in a complex biochemical process called melanogenesis, which is influenced by extrinsic and intrinsic factors, such as hormones, inflammation, age, and ultraviolet light exposure. We describe three methods to determine the hypopigmentation activity of chemicals or natural substances in melanocytes: measurement of the 1) cellular tyrosinase activity and 2) melanin content, and 3) staining and quantifying cellular melanin with image analysis.
In melanogenesis, tyrosinase catalyzes the rate-limiting step that converts L-tyrosine into 3,4-dihydroxyphenylalanine (L-DOPA) and then into dopaquinone. Therefore, the inhibition of tyrosinase is a primary hypopigmentation mechanism. In cultured melanocytes, tyrosinase activity can be quantified by adding L-DOPA as a substrate and measuring dopaquinone production by spectrophotometry. Melanogenesis can also be measured by quantifying the melanin content. The melanin-containing cellular fraction is extracted with NaOH and melanin is quantified spectrophotometrically. Finally, the melanin content can be quantified by image analysis following Fontana-Masson staining of melanin. Although the results of these in vitro assays may not always be reproduced in human skin, these methods are widely used in melanogenesis research, especially as the initial step to identify potential hypopigmentation activity. These methods can also be used to assess melanocyte activity, growth, and differentiation. Consistent results with the three different methods ensure the validity of the effects.

We describe three experimental methods for evaluating the hypopigmentation activity of chemicals in vitro: quantification of 1) cellular tyrosinase activity and 2) melanin content, and 3) measurement of melanin by cellular melanin staining and image analysis.

This study presents laboratory methods for the quantification of hypopigmentation activity in vitro. Melanin, the major pigment in melanocytes, is ...

Isolation of Physiologically Active Thylakoids and Their Use in Energy-Dependent Protein Transport Assays

Chloroplasts are the organelles in green plants responsible for carrying out numerous essential metabolic pathways, most notably photosynthesis. Within the chloroplasts, the thylakoid membrane system houses all the photosynthetic pigments, reaction center complexes, and most of the electron carriers, and is responsible for light-dependent ATP synthesis. Over 90% of chloroplast proteins are encoded in the nucleus, translated in the cytosol, and subsequently imported into the chloroplast. Further protein transport into or across the thylakoid membrane utilizes one of four translocation pathways. Here, we describe a high-yield method for isolation of transport-competent thylakoids from peas (Pisum sativum), along with transport assays through the three energy-dependent cpTat, cpSec1, and cpSRP-mediated pathways. These methods enable experiments relating to thylakoid protein localization, transport energetics, and the mechanisms of protein translocation across biological membranes.

We present protocols herein for high-yield isolation of physiologically active thylakoids and protein transport assays for the chloroplast twin arginine translocation (cpTat), secretory (cpSec1), and signal recognition particle (cpSRP) pathways.

Chloroplasts are the organelles in green plants responsible for carrying out numerous essential metabolic pathways, most notably photosynthesis. Within ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

In Vitro Ubiquitination and Deubiquitination Assays of Nucleosomal Histones

Ubiquitination is a post-translational modification that plays important roles in various signaling pathways and is notably involved in the coordination of chromatin function and DNA-associated processes. This modification involves a sequential action of several enzymes including E1 ubiquitin-activating, E2 ubiquitin-conjugating and E3 ubiquitin-ligase and is reversed by deubiquitinases (DUBs). Ubiquitination induces degradation of proteins or alteration of protein function including modulation of enzymatic activity, protein-protein interaction and subcellular localization. A critical step in demonstrating protein ubiquitination or deubiquitination is to perform in vitro reactions with purified components. Effective ubiquitination and deubiquitination reactions could be greatly impacted by the different components used, enzyme co-factors, buffer conditions, and the nature of the substrate.&#160; Here, we provide step-by-step protocols for conducting ubiquitination and deubiquitination reactions. We illustrate these reactions using minimal components of the mouse Polycomb Repressive Complex 1 (PRC1), BMI1, and RING1B, an E3 ubiquitin ligase that monoubiquitinates histone H2A on lysine 119. Deubiquitination of nucleosomal H2A is performed using a minimal Polycomb Repressive Deubiquitinase (PR-DUB) complex formed by the human deubiquitinase BAP1 and the DEUBiquitinase ADaptor (DEUBAD) domain of its co-factor ASXL2. These ubiquitination/deubiquitination assays can be conducted in the context of either recombinant nucleosomes reconstituted with bacteria-purified proteins or native nucleosomes purified from mammalian cells. We highlight the intricacies that can have a significant impact on these reactions and we propose that the general principles of these protocols can be swiftly adapted to other E3 ubiquitin ligases and deubiquitinases.

Ubiquitination is a post-translational modification that plays important roles in cellular processes and is tightly coordinated by deubiquitination. Defects in both reactions underlie human pathologies. We provide protocols for conducting ubiquitination and deubiquitination reaction in vitro using purified components.

Ubiquitination is a post-translational modification that plays important roles in various signaling pathways and is notably involved in the coordination ...

A Planarian Motility Assay to Gauge the Biomodulating Properties of Natural Products

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and withdrawal properties of natural products is described. Experimental assays benefitting from unique aspects of planarian physiology have been applied to studies on wound healing, regeneration, and tumorigenesis. In addition, because planarians exhibit sensitivity to a variety of environmental stimuli and are capable of learning and developing conditioned responses, they can be used in behavioral studies examining learning and memory. Planarians possess a basic bilateral symmetry and a central nervous system that uses neurotransmitter systems amenable to studies examining the effects of neuromuscular biomodulators. Consequently, experimental systems monitoring planarian movement and motility have been developed to examine substance addiction and withdrawal. Because planarian motility offers the potential for a sensitive, easily standardized motility assay system to monitor the effect of stimuli, the planarian locomotor velocity (pLmV) test was adapted to monitor both stimulation and withdrawal behaviors by planarians through the determination of the number of grid lines crossed by the animals with time. Here, the technique and its application are demonstrated and explained.

Planarian motility is used to gauge the stimulant and withdrawal properties of natural products when compared to the movement of the animals in spring water alone.

A straightforward, controllable means of using the non-parasitic planarian, Dugesia tigrina, a free-living aquatic flatworm, to study the stimulant and ...

Time-resolved F&#246;rster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human Cells

The Janus kinase (JAK)/signal transducer and activator of transcription (STAT) signaling pathway plays a crucial role in mediating cellular responses to cytokines and growth factors. STAT proteins are activated by tyrosine phosphorylation mediated mainly by JAKs. The abnormal activation of STAT signaling pathways is implicated in many human diseases, especially cancer and immune-related conditions. Therefore, the ability to monitor STAT protein phosphorylation within the native cell signaling environment is important for both academic and drug discovery research. The traditional assay formats available to quantify phosphorylated STAT proteins include western blotting and the enzyme-linked immunosorbent assay (ELISA). These heterogeneous methods are labor-intensive, low-throughput, and often not reliable (specific) in the case of western blotting. Homogeneous (no-wash) methods are available but remain expensive. 
Here, detailed protocols are provided for the sensitive, robust, and cost-effective measurement in a 384-well format of endogenous levels of phosphorylated STAT1 (Y701), STAT3 (Y705), STAT4 (Y693), STAT5 (Y694/Y699), and STAT6 (Y641) in cell lysates from adherent or suspension cells using the novel THUNDER time-resolved F&#246;rster resonance energy transfer (TR-FRET) platform. The workflow for the cellular assay is simple, fast, and designed for high-throughput screening (HTS). The assay protocol is flexible, uses a low-volume sample (15 &#181;L), requires only one reagent addition step, and can be adapted to low-throughput and high-throughput applications. Each phospho-STAT sandwich immunoassay is validated under optimized conditions with known agonists and inhibitors and generates the expected pharmacology and Z'-factor values. As TR-FRET assays are ratiometric and require no washing steps, they provide much better reproducibility than traditional approaches. Together, this suite of assays provides new cost-effective tools for a more comprehensive analysis of specific phosphorylated STAT proteins following cell treatment and the screening and characterization of specific and selective modulators of the JAK/STAT signaling pathway.

Time-resolved F&#246;rster resonance energy transfer cell-based assay protocols are described for the simple, specific, sensitive, and robust quantification of endogenous phosphorylated signal transducer and activator of transcription (STAT) 1/3/4/5/6 proteins in cell lysates in a 384-well format.

The Janus kinase (JAK)/signal transducer and activator of transcription (STAT) signaling pathway plays a crucial role in mediating cellular responses to ...

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) establish and maintain the DNA methylation pattern. Changes to the normal DNA methylation pattern are linked to cancer development and progression, making DNMTs potential cancer drug targets. Thus, identifying and characterizing novel small molecule inhibitors of these enzymes is of great importance. This paper presents a protocol that can be used to screen for DNA methyltransferase inhibitors. The continuous coupled kinetics assay allows for initial velocities of DNA methylation to be determined in the presence and absence of potential small molecule inhibitors. The assay uses the methyl-sensitive endonuclease Gla I to couple methylation of a hemimethylated DNA substrate to fluorescence generation.
This continuous assay allows for enzyme activity to be monitored in real time. Conducting the assay in small volumes in microtiter plates reduces the cost of reagents. Using this assay, a small example screen was conducted for inhibitors of DNMT1, the most abundant DNMT isozyme in humans. The highly substituted anthraquinone natural product, laccaic acid A, is a potent, DNA-competitive inhibitor of DNMT1. Here, we examine three potential small molecule inhibitors &#8212; anthraquinones or anthraquinone-like molecules with one to three substituents &#8212; at two concentrations to describe the assay protocol. Initial velocities are used to calculate the percent activity observed in the presence of each molecule. One of three compounds examined exhibits concentration-dependent inhibition of DNMT1 activity, indicating that it is a potential inhibitor of DNMT1.

DNA methyltransferases are potential cancer drug targets. Here, a protocol is presented to assess small molecules for DNA methyltransferase inhibition. This assay utilizes an endonuclease to couple DNA methylation to fluorescence generation and allows for enzyme activity to be monitored in real time.&#160;

DNA methylation, a form of epigenetic gene regulation, is important for normal cellular function. In cells, proteins called DNA methyltransferases (DNMTs) ...

Myosin-Specific Adaptations of In vitro Fluorescence Microscopy-Based Motility Assays

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Time-resolved Förster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human Cells

Continuous Fluorescence-Based Endonuclease-Coupled DNA Methylation Assay to Screen for DNA Methyltransferase Inhibitors