1024

Biophysical Journal

Volume 100

February 2011

1024–1033

Evidence for Pre- and Post-Power Stroke of Cross-Bridges of Contracting
Skeletal Myofibrils
K. Midde,†‡ R. Luchowski,†‡k H. K. Das,§{ J. Fedorick,†‡ V. Dumka,†‡ I. Gryczynski,†‡ Z. Gryczynski,†‡and J. Borejdo†‡*†Department of Molecular Biology and Immunology, ‡Center for Commercialization of Fluorescence Technologies,
Department of Pharmacology and Neuroscience, and {Institute of Cancer Research, University of North Texas Health Science Center,
Fort Worth, Texas; and kDepartment of Physics, Maria Curie-Sklodowska University, Lublin, Poland§

ABSTRACT We examined the orientational fluctuations of a small number of myosin molecules (approximately three) inworking skeletal muscle myofibrils. Myosin light chain 1 (LC1) was labeled with a fluorescent dye and exchanged with the native
LC1 of skeletal muscle myofibrils cross-linked with 1-ethyl-3-[3(dimethylamino) propyl] carbodiimide to prevent shortening. We
observed a small volume within the A-band (~1015 L) by confocal microscopy, and measured cyclic fluctuations in theorientation of the myosin neck (containing LC1) by recording the parallel and perpendicular components of fluorescent lightemitted by the fluorescently labeled myosin LC1. Histograms of orientational fluctuations from fluorescent molecules in rigor
were represented by a single Gaussian distribution. In contrast, histograms from contracting muscles were best fit by at least
two Gaussians. These results provide direct evidence that cross-bridges in working skeletal muscle assume two distinct conformations, presumably corresponding to the pre- and post-power-stroke states.

INTRODUCTION
Muscle contraction results from the cyclic interaction of
myosin heads with actin filaments. A critical intermediatestep in this cycle is the power stroke, a conformationalchange in myosin that causes force production and/or slidingof myosin-containing filaments relative to actin-containing
filaments. The biochemistry of the actomyosin interaction
is well understood (1–4) and implies that the formation ofa pre-power-stroke state results from the binding of
myosin-containing hydrolytic products to actin. The releaseof phosphate causes a force-generating conformationalchange of the bound head that leads to the formation ofa post-power-stroke state. In this state, both ADP and actin
are bound to a myosin head (ADP is bound weakly, hence
the term ‘‘weak ADP binding state’’ (5)). Dissociation of
ADP transforms this state to a rigor conformation in which
myosin, devoid of hydrolytic products, binds strongly toactin. The conformation of a cross-bridge in a weak ADP
binding state is nearly the same as in rigor. After that, thecross-bridge enters the so-called near-rigor state, in whicha head is still bound to actin but allows ATP binding. Releaseof ADP causes the head to bind to a new ATP molecule andthus detach from actin and hydrolyze ATP (6). The prepower-stroke and post-power-stroke states are also known
as closed and open states, respectively (1). In this work we
examine the distribution of cross-bridges between the preand post-power-stroke states of the myosin ATPase cycle.
Strong evidence for the existence of distinct pre- and postpower-stroke states has been provided by in vitro singlemolecule (7), crystallography (5), and electron microscopy
Submitted August 18, 2010, and accepted for publication January 5, 2011.
*Correspondence: Julian.Borejdo@unthsc.edu(8) studies of the processive myosin V molecule. Earlier,elegant EPR studies (9) suggested the existence of threestructural states of myosin II in working muscle fibers. One
state, the rigor state, has been well characterized in ex vivo
muscle. This conformation has been extensively described,
because all of the cross-bridges in rigor are immobile andassume a well-defined orientation with respect to the muscle
axis (10–12). Investigators have obtained valuable information about cross-bridge orientation, number, position, andangle in actin target zones by measuring large assemblies
of cross-bridges in insect flight (13) and vertebrate skeletal
muscle (9,12,14–17). In this work, we are specifically interested in the distribution of cross-bridge orientations duringcontraction. Because cross-bridges act asynchronously
during contraction, the resulting information about spatial
distribution is not very useful if it is averaged over many
molecules. Therefore, it is necessary to examine a distributionof cross-bridge fluctuations about the average value, i.e., tofollow the motion of mesoscopic (affected by fluctuations
around the average (18)) of myosin molecules. The size ofthe fluctuation is inversely proportional to the number ofmolecules under study, and therefore we looked at very small
assemblies of cross-bridges (three to four molecules) in
isometrically contracting skeletal muscle myofibrils.
The method applied in this study is based on the ability toobserve a few molecules of myosin in an ex vivo muscle, as
illustrated in Fig. 1. Muscle myosin light chain 1 (LC1) islabeled with rhodamine and exchanged with native LC1 of
a myofibril. A small volume within labeled muscle(1016 L) is illuminated by a focused laser beam (Fig. 1,
green hourglass), and the fluorescence is observed by
confocal microscopy. The detection volume (DV; dashedline) of the microscope is an ellipsoid of revolution whose

Editor: Christopher Lewis Berger.
Ó 2011 by the Biophysical Society
0006-3495/11/02/1024/10 $2.00

doi: 10.1016/j.bpj.2011.01.007

Pre- and Post-Power Stroke

1025

MATERIALS AND METHODS
Chemicals and solutions
Tetramethylrhodamine-5-iodoacetamide dihydroiodide (5-TMRIA, single
isomer) was purchased from Molecular Probes (Eugene, OR; Cat. No.
T-6006). All other chemicals were obtained from Sigma-Aldrich (St. Louis,
MO). The composition of the solutions was identical to that used by Muthu
et al. (26).

LC1 expression

FIGURE 1 Origin of fluorescence fluctuations. A small fraction of
myosin within the myofibril is bound to rhodamine-labeled LC1 (redcircle). In rigor (top panel) there are no fluctuations in the orientation of
the fluorophore’s transition dipole (red arrow). During contraction (bottom
panel) the myosin cross-bridges are cycling, which changes the orientationof the rhodamine’s transition dipole and causes fluctuations of polarized
intensity. We measure fluctuations in dipole orientation by recording
parallel (k) and perpendicular (t) components of the emitted fluorescentintensity by the cross-bridge-bound fluorophores.

small diagonal measures 1.2 mm. If the degree of labeling issufficiently small, this volume will contain three to four fluorescent molecules of myosin (see the ‘‘Number of observed
molecules’’ section in the Results). In rigor muscle (toppanel), the myosin cross-bridges are stationary and no fluctuations in the orientation of the transition dipole occur. Duringcontraction, on the other hand, the myosin cross-bridges
undergo power-stroke cycles (bottom panel). The powerstroke changes the orientation of a transition dipole of rhodamine attached to LC1, i.e., the orientation of rhodamine dipole
fluctuates in time. We measured these fluctuations by simultaneously recording parallel (Ik) and perpendicular (It) components of fluorescent light emitted by a myosin-boundfluorophore. Each of these components reflects orientationalchanges of the myosin head C-terminus. The normalized
difference between these components, i.e., the polarized fluorescence (P), is a sensitive indicator of the orientation of thetransition dipole of the fluorophore (15,19–25). We performed
an elementary statistical analysis to analyze the observed fluctuations, and found that the fluctuations of the rigor muscles
were distributed according to a single Gaussian distribution.
In contrast, the histograms of contracting muscles could be
fitted by at least two Gaussians. This provides evidence thatcross-bridges in working skeletal muscle assume two distinct
conformations, presumably corresponding to the pre-and
post-power stroke states of the myosin ATPase cycle.

A pQE60 vector containing recombinant LC1 with a single cysteine residue(Cys-178) was donated by Dr. Susan Lowey (University of Vermont). The
plasmid DNA was transformed into Escherichia coli M15 competent cells,
and recombinant clones were selected by ampicillin resistance. The
LC1-cDNA insert of the clones was confirmed by DNA sequencing ofboth strands (Iowa State University of Science and Technology). LC1 protein
was overexpressed in Luria broth containing 100 mg/ml of ampicillin by
induction with IPTG. His-tagged LC1 protein was affinity-purified ona Ni-NTA column according to the manufacturer’s protocol. The imidazole-eluted fractions were run on SDS-PAGE followed by Western analysis
with Anti-LCN1 antibodies (Abcam, Cambridge, MA). Fractions containing
LC1 were pooled together and dialyzed with buffer (50 mM KCl and 10 mM
phosphate buffer, pH 7.0). Dialyzed protein showed a single ~25-kDa bandon SDS-PAGE after Coomassie staining (see Fig. S1 in the Supporting Material). Protein concentration was determined by using the Bradford assay.

LC1 labeling
Purified LC1 was dialyzed against buffer A (50 mM KCL and 10 mM phosphate buffer, pH 7.0) and fluorescently labeled by incubation with a 5 molar
excess of 5-TMRIA for 6 h in buffer A on ice. To eliminate unbound dye,the solution was passed through a Sephadex G50 column.

Degree of labeling of LC1
The concentrations of LC1 protein and bound 5-TMRIA were determined toestimate the degree of labeling. Protein concentration was determined by
the Bradford assay, and 5-TMRIA concentration was determined fromthe peak absorbance (3 ¼ 87,000 M1cm1 at 535 nm) obtained on a Varian
Eclipse spectrometer (Varian, Palo Alto, CA). The concentrations of theprotein and 5-TMRIA dye were found to be the same, indicating that thedye and protein were bound in a 1:1 ratio.

Preparation of myofibrils
Rabbit psoas muscle bundles were washed with an ice-cold EDTA-rigor
solution (50 mM KCl, 5 mM EDTA, 10 mM TRIS-HCl, pH 7.5) for 0.5 h,followed by an extensive washing with Mg2þ-rigor solution (50 mM KCl,
2 mM MgCl2, 10 mM TRIS-HCl, pH 7.5) and then with Ca2þ-rigor solution(50 mM KCl, 0.1 mM CaCl2, 10 mM TRIS-HCl, pH 7.5). The fiber bundle
was then homogenized with the use of a Heidolph Silent Crusher S homogenizer for 20 s (with a break to cool after 10 s) in Mg2þ-rigor solution.

LC1 exchange into myofibrils
Fluorescently tagged LC1 (R-LC1) was incubated with 1 mg/ml of freshly
prepared myofibrils in light-chain exchange solution (15 mM KCl, 5 mM
EDTA, 5 mM DTT, 10 mM KH2PO4, 5 mM ATP, 1 mM TFP, and10 mM imidazole, pH 7) for 5 min at 30 C (12). The concentration of
R-LC1 was 3 nM unless otherwise specified.
Biophysical Journal 100(4) 1024–1033

1026

Probability distribution measurements
Fluorescence fluctuations were acquired on an Alba-FCS confocal system(ISS, Urbana, IL) coupled to an Olympus IX 71 microscope. The experimental arrangement was the same as that previously described by Borejdo
et al. (27). In brief, excitation was provided by a 532 nm CW laser. A polarizer
was inserted before the entrance to the microscope to ensure that the exciting
light was linearly polarized, and the resulting laser polarization was verticalwith respect to the microscope stage. The confocal pinhole was 50 mm. The
emitted fluorescent light was split by a prism, and each component was
detected by a separate Avalanche PhotoDiode (APD) through k and toriented analyzers. Myofibrils were always placed with their long axis aligned
vertically on the microscope stage. We collected 2 M data points every 10 ms
for 20 s and smoothed the data by binning 1000 points. Before each experiment, the instrument was calibrated and optimized with a 50 nM solutionof rhodamine G. Following the notation of Tregear and Mendelson (22), let
kIt be the polarized intensity obtained with the exciting and detected lightpolarized parallel to the myofibril axis, and kIt be the polarized intensity
obtained with exciting and detected light polarized parallel and perpendicular
to the myofibril axis, respectively. Channel 1 (ch1) was used to detect kIt,
and channel 2 (ch2) was used to detect kIk. We calculated polarization as(kIt  kIk)/(kIt þ kIk). Thus, the reported polarization values are negativevalues of the conventional Pk polarization parameter.

Steady-state force measurements
A small bundle of skinned skeletal muscle fibers (two to five) from rabbit
psoas were attached by tweezer clips to a force transducer (Scientific Instruments, Heidelberg, Germany). The fibers were stretched until taut in 1 ml of
pCa 5 rigor solution (50 mM KCl, 0.1 mM CaCl2, 10 mM TRIS-HCl,
pH 7.5), and then a contracting solution (50 mM KCl, 0.1 mM CaCl2,
5 mM MgATP, 10 mM TRIS-HCl, pH 7.5) was added to test for maximal
steady-state force. We tested whether LC1 exchange modified the tension of
the muscle fibers. Fibers were exchanged with 100 nM fluorescent LC1 at
30 C for different periods of time. In N ¼ 60 experiments on different
fibers, the average maximal force per unlabeled fiber was 1.6 5 0.1 mN(mean 5 SE). The force was also measured after exchange for 5 (N ¼
6), 10 (N ¼ 5), 15 (N ¼ 8), or 30 (N ¼ 6) min. At 5 and 10 min after
exchange, the force was unchanged. At 15 min, the force was actually
increased to 1.9 mN/fiber and again decreased to the unlabeled value of
1.6 mN/fiber at 30 min. Because in our experiments the myofibils were
exchanged for 5 min with 3 nM fluorescent LC1, we conclude that labelinghas no effect on fiber tension or myofibril function.

Midde et al.
Germany). The excitation was provided by a 475-nm pulsed diode laser,
and the observation was conducted through a 590 nm monochromatorwith a supporting 590-nm long-pass filter. The full width at half-maximumof the pulse response function was 68 ps (measured by PicoQuant), and thetime resolution was better than 10 ps. We analyzed the intensity decays interms of a multiexponential model using FluoFit software (PicoQuant). All
experiments were performed at room temperature (~23 C).

Statistical analysis
Statistical analysis was performed with the use of SigmaPlot 11.02 andPeakFit v. 4.12 (Systat, San Jose, CA) and Origin v. 8.5 (OriginLab, Northampton, MA) software. The quality of fit to either a single or double
Gaussian function was assessed by the standard error (PeakFit, San Jose,CA) or reduced c2 (Origin) parameters.

RESULTS
Imaging
A typical fluorescence lifetime image of a rigor myofibril isshown in Fig. 2. All A-bands were fluorescently labeled.
The red circle in Fig. 2 A indicates a two-dimensional
projection of the confocal aperture on the image plane,
and its diameter (1.2 mm) is equal to the diameter of theconfocal aperture (50 mm) divided by the magnification of
the objective (40). Images are shown at various degrees

Cross-linking
In the presence of Ca2þ and ATP, the myofibrils shortened, which made it
impossible to record polarized intensities during contraction. Therefore, weprevented the myofibrils from shortening by cross-linking them with thewater-soluble cross-linker 1-ethyl-3-[3-(dimethylamino)propyl]carbodiimide (EDC) (28,29). Myofibrils (1 mg/ml) in rigor solution (50 mM KCl,
20 mM EDC, 0.1 mM CaCl2, 10 mM TRIS-HCl, pH 7.5) were incubatedfor 20 min at room temperature. The reaction was stopped by addition of
20 mM DTT. The pH of the solution (7.4) remained unchanged throughout
the 20 min reaction. We verified the lack of shortening by imaging themyofibrils by differential contrast microscopy, and by fluorescence microscopy after labeling the myofibrils with 10 nM rhodamine-phalloidin (30).

Time-resolved anisotropy and lifetime
measurements
We measured the fluorescence anisotropy and lifetimes using a timedomain technique and a FluoTime 200 fluorometer (PicoQuant, Berlin,
Biophysical Journal 100(4) 1024–1033

FIGURE 2 Lifetime images of a rigor myofibril. The color bar to theright of image A is the lifetime scale from 0 (blue) to 6 (red) ns. The redcircle is a projection of the confocal aperture on the sample plane (diameter
1.2 mm). Native LC1 molecules were exchanged in myofibrils with 4 nM(A), 2 nM (B), 1.2 nM (C), and 0.8 nM (D) of R-LC1. Sarcomere
length ¼ 3.1 mm (A), 3.0 mm (B), 3.0 mm (C), and 2.9 mm (D). Images
were acquired on a PicoQuant Micro Time 200 confocal lifetime microscope. The sample was excited with a 470 nm pulse of light and the emission was collected on an APD through a LP500 filter.

Pre- and Post-Power Strokeof labeling. Native LC1 of the myofibrils was exchanged
with 4 nM (A), 2 nM (B), 1.2 nM (C), and 0.8 nM (D) of
R-LC1. In the experiments reported here, the myofibrils
were exchanged with 3 nM R-LC1.
LC1 is immobilized by the myosin neckWe measured the decay of anisotropy of R-LC1 exchanged
into myofibrils to test whether rhodamine was rigidly immobilized on the surface of LC1. This was done to ensure thatthe orientation of the transition dipole of the fluorophore
would reflect the orientation of the neck portion of the
myosin head. Anisotropy is defined as r ¼ (Ik  It)/(Ik þ
2It). (The decay is shown in Fig. 4.) The decay of free
TMRIA was best fit by a double exponential curve r(t) ¼
Roþ a  exp(t/q1) þ b  exp(t/q2), where Ro ¼ 0.05
is the value of anisotropy at infinite time, and q1 ¼ 0.3
and q2 ¼ 608.7 ns are the rotational correlation times(Fig. 3 A). The fast decay accounted for most of the anisotropy decay (93.1%), and thus the slow correlation time accounted for only 6.9% of the decay, probably due to

FIGURE 3 Anisotropy decay of 0.1 mM free TMRIA (A) and 50 mM
myofibrils (B) after exchange with 2.5 mM R-LC1 into 1 mg/ml myofibrils
for 5 min at 30 C.

1027

aggregates of rhodamine. The decay of R-LC1 bound to
myofibrillar myosin was best fit by the same double exponential function, but this time Ro ¼ 0.33, q1 ¼ 67.7, andq2 ¼ 0.9 ns (Fig. 3 B). The slow and fast correlation times
accounted for 82.1% and 17.9%, respectively, of the decay.
The short and long correlation times were most likely due tothe rotation of the rhodamine moiety on LC1 and the rotation of bound LC1, respectively. The maximum value ofanisotropy was 0.384, and this high value of initial anisotropy indicates that the absorption and emission dipoles of
rhodamine are nearly parallel.
Number of observed molecules
Because asynchronous operation of a large number of crossbridges will average out any fluctuations, it is essential to
make sure that the data are collected from only a few molecules. Therefore, we labeled the myosin very sparsely withfluorescent LC1 (myofibrils were incubated with 3 nM
rhodamine-LC1) and then placed a myofibril in the focusof the microscope. To determine how many molecules
contributed to the observed fluorescence, we measured the
autocorrelation function (ACF) of fluorescence fluctuations
for selected fluorophore concentrations. The value of the
ACF at delay time 0 [G(0)] is equal to the inverse of thenumber of molecules N contributing to the signal, N ¼ 1/
G(0) (31,32). The ACFs were obtained for solutions of
TMRIA in the range of 1.7–8.5 nM. An example of the
ACF obtained at a concentration of 6.8 nM is shown in
Fig. 4 A. The value of G(0) was 0.07, giving an average
number of molecules of ~14. The number of molecules estimated from the cross-correlation functions of ch1 and ch2
are plotted in Fig. 4 B as red and green circles, respectively.
Extrapolation yields the approximate number of photons per
one fluorophore as ~1400 counts/s per channel.
We were unable to obtain the ACFs at very low concentrations of the dye (%3.4 nM TMRIA) because the ACFs
were simply too noisy. Therefore, we measured the intensityof the signal as a function of the concentration of the dye.
The plot of concentration versus counts in ch1 and ch2 isshown in Fig. 4 C. The DV of the microscope is an ellipsoidof revolution whose waist (1.2 mm) is equal to the diameterof the confocal pinhole (2uo ¼ 50 mm) divided by themagnification of the objective (40). The ellipsoid isassumed to have a waist uo and height, zo, equal to thethickness (1 mm) of a typical myofibril. Therefore, DV ¼
4/3puo2zo is ~1 mm3. This is approximately equal to thevolume of a typical half-sarcomere. Knowing the DV andconcentration, we calculated the number of molecules inthe DV. The concentration of 1.7 nM TMRIA corresponds
to a single fluorophore. Extrapolation yields the approximate number of photons per one fluorophore: ~100
counts/s per channel. Averaging extrapolated values from
Fig. 4, B and C, yields ~750 counts/s from a single
fluorophore.
Biophysical Journal 100(4) 1024–1033

1028

Midde et al.

laser light was also vertical, i.e., parallel to the myofibrillar
axis. The fluorescence emanating from the DV was split
50/50 by a beam splitter and projected through 50 mm
confocal apertures onto the photosensitive surface of the
two APDs. Fig. S2 shows a typical time course of polarized
intensity of a rigor (A) and contracting (B) myofibril. The
original data were collected every 10 ms, but to smoothdata and to keep the files to a manageable size, we binned
1000 points together to obtain a time resolution of 10 ms.
The vertical scale is the number of counts during 10 ms.
Estimation of the number of molecules
contributing to the signal from the myofibril
The average intensities from contracting myofibrils in ch1
and ch2 were 21 and 23, respectively (Fig. S2). For rigor
myofibrils, the averages were 23 and 27 counts/10 ms,
respectively (note that Fig. S2 is a bar plot). Thus, theaverage (23 counts/10 ms) corresponds to 2300/750, or~3 myosin molecules.
Skewness and kurtosis

FIGURE 4 (A) Cross-correlation function of 6.8 nM TMRIA. The
number of molecules in the DV was obtained from the inverse of thecross-correlation value extrapolated at a delay time equal to zero. (B)Plot of the number of molecules in the DV obtained from the inverse ofcross-correlation function versus signal intensity for ch1 (red dots) andch2 (green dots). The solid line is a single exponential, three-parameterfit (y ¼ yo þ abx). (C) Signal intensity plotted as a function of TMRIA
concentration (1.7–8.5 nM) for ch1 (red dots) and ch2 (green dots).
The solid line is a single exponential, three-parameter fit (y ¼ yo þ abx).
The number of molecules in the DV at a TMRIA concentration of
1.7 nM corresponds to one molecule, and the number of photons per fluorophore in plots B and C was estimated to be ~750 counts/s per channel.

Intensity fluctuations during rigor andcontraction of muscle myofibrils
A myofibril was placed on the stage of the confocal microscope with the long axis oriented vertically on the microscope stage. The linear polarization of the illuminating
Biophysical Journal 100(4) 1024–1033

Fig. 5 shows examples of six histograms of polarized fluorescence randomly selected from a pool of 20 histograms
obtained from myofibrils in rigor. A histogram can be quantitatively characterized by the values of kurtosis and skewness. A positive skewness means that the tail of the curveis directed toward positive values of the histogram relative
to a Gaussian distribution. This was the case for both rigor
and contraction (see below). A positive kurtosis is expressedby long tails and higher peaks compared with the Gaussian
curves. This was the case for rigor myofibrils. A negative
kurtosis means that the tails are smaller than those ofGaussian curves. This was the case for contracting myofibrils (see below). Zero skewness and kurtosis means thatthe histograms are Gaussian. This was the case for theuncross-linked myofibrils in rigor. Table 1 shows the valuesof skewness and kurtosis. The differences in both skewness
and kurtosis between EDC-rigor and contracting myofibrils
were both statistically significant. The average values ofskewness for rigor histograms were influenced by crosslinking. The difference was statistically significant.
Effect of EDC
We sought to ascertain whether cross-linking makes a difference in the rigor distribution. Fig. S3 of the shows a rigor
distribution obtained from uncross-linked myofibrils. The
goodness of fit, assessed by c2, showed that none of thecurves were significantly improved by the addition of
a second Gaussian. The second peak accounts for (clockwise, starting from upper left) 0, 6.4, 0, 0.3, 0, and 8.3%of the total signal (average ¼ 2.5%). The average waseven smaller in the remaining 12 experiments. Even thoughPre- and Post-Power Stroke

1029

FIGURE 5 Examples of histograms of polarized fluorescence from rigor (top panel) and relaxed (bottom panel) myofibrils. Horizontal scale: The negativevalue of parallel fluorescence polarization. Vertical scale: The number of events during 20 s. The position of peaks for rigor myofibrils were (top panel,clockwise from the top left): P ¼ 0.07, P ¼ 0.05, P ¼ 0.08, P ¼ 0.02, P ¼ 0.00, P ¼ 0.07. The blue line shows the fit to a single Gaussian distribution.
For relaxed myofibrils (bottom panel), the parameters (area, center, width, and height) for each peak of a two Gaussian fit are given in clockwise order, starting from the upper left. Upper left: minor peak (red) 0.85, 0.18, 0.10, 6.82; major peak (green) 14.89, 0.03, 0.21, 57.01. Upper middle: minor peak (red)0.64, 0.03, 0.00, 115.88; major peak (green) 17.16, 0.02, 0.22, 61.66. Upper right: minor peak (red) 0.11, 0.11, 0, 86.96; major peak (green) 12.63, 0.07,
0.19, 51.74. Lower left: minor peak (red) 1.09, 0.05, 0.09, 10.09; major peak (green) 12.10, 0.09, 0.18, 53.36. Lower middle: minor peak (red) 0.23, 0.05,
0.02, 9.54; major peak (green) 14.24, 0.07, 0.21, 53.62. Lower right: minor peak (red) 0.47, 0.01, 0.05, 7.57; major peak (green) 14.30, 0.00, 0.23, 48.84.
The contribution of the minor peak (red line) makes little difference relative to the second peak (green line) of a double Gaussian distribution fit. The blue lineis the sum of red and green line fits.

the cross-linking is extensive (29), we believe that it makes
no difference in terms of our conclusions in this study.
Cross-linking occurs between myosin heads and actin, andbetween LMM tails, which are remote enough from LC1
to have no effect on the observed fluctuations.
Rigor histograms
The fit of representative histograms in rigor with a threeparameter Gaussian curve (y ¼ a exp[0.5(x  xo/b)2]) is
shown in the top panel of Fig. 5 (blue line). The goodnessof fit, assessed by standard error in the PeakFit program,showed that none of the curves were significantly improvedby the addition of a second Gaussian. This point is illustrated in Fig. S4, which shows the two-Gaussian fit to thesame data. The ratios of the areas under the major (green)and minor (red) peaks were (clockwise from upper left)
17.5, 18.1, 11.6, 59.5, 12.6, and 55.0, with an average of
29.0, indicating that adding the second Gaussian made on
average only a 3.4% difference. The average position ofa single peak was P ¼ 0.05 5 0.08 (in an oriented, immobile system, the fact that the polarization ¼ 0 does notnecessarily mean that the dipoles are randomly arranged).
Because the absorption and emission dipoles of rhodamine
are nearly parallel (time zero anisotropy z 0.4; Fig. 3)
and the myofibrillar axis is parallel to the direction ofBiophysical Journal 100(4) 1024–1033

1030

Midde et al.

TABLE 1 Comparison of kurtosis and skewness parametersfrom fluorescence polarization fluctuation histograms of rigor
and contracting skeletal myofibrils
Myofibrils
Rigor
EDC-rigor
Relaxation
EDC-relaxation
Contraction

Skewness

KurtosisN0.02 5 0.04
1.12 5 0.34
1.04 5 0.19
1.48 5 0.54
0.86 5 0.16

0.06 5 0.18
0.84 5 1.63
0.20 5 0.81
3.00 5 3.05
0.59 5 0.27

18
18*
18
18
18*

Errors are given as SD, and N is the number of experiments.
*Two histograms with extreme maxima and minima were rejected.

polarization of exciting light, this indicates that the transition dipoles of rhodamine in rigor are oriented mostly
perpendicular to myofibrillar axis.
Relaxation histograms
Histograms of relaxed myofibrils were also best fit by a single
Gaussian. The bottom panel of Fig. 5 shows the representative histograms during relaxation. The ratios of the areas
under the major (green) and minor (red) peaks were (clockwise from upper left) 17.4, 26.8, 114.5, 12.0, 61.7, and31.1, with an average of 43.9, indicating that adding thesecond Gaussian made on average only a 2.2% difference.
Contraction histograms
The histograms of polarized fluorescence fluctuations obtained from contracting myofibrils were significantly
different from those observed in rigor and relaxation. Fig. 6

shows examples of six histograms of polarized fluorescencerandomly selected from a pool of 20 histograms of contracting myofibrils. The average skewness and kurtosis were0.86 5 0.16 (SD) and 0.59 5 0.27 (SD), respectively.
Both skewness and kurtosis were statistically significantly
different during rigor and contraction at a level of 0.69%
for skewness (t ¼ 2.87, P ¼ 0.0069, 34 degrees of freedom)
and at a level of <0.07% for kurtosis (t ¼ 3.68,
P ¼ 0.00078, 34 degrees of freedom), respectively. The standard error test of the PeakFit program showed that 14 of 20
histograms were best fit by two Gaussians. The average positions of the peaks were P ¼ 0.14 5 0.04 for peak 1 andP ¼ 0.31 5 0.05 for peak 2 (Table 2). The positions of
the two peaks were significantly statistically different at
P < 0.001% (t ¼ 9.92). Most of the power was contained inpeak 1, although on two occasions the peak 2 was larger
than peak 1 (e.g., Fig. 6, right upper corner). The average
ratio, i.e., the relative contribution of both Gaussians, was
5.7 5 7.7. The overall experimental curve (blue) is thesum of the red (peak 1) and green (peak 2) Gaussians.
Reversibility
To ensure that the second Gaussian peak observed duringcontraction was not an artifact induced by the contraction
protocol itself, we examined the orientational distributions
in rigor before and after contraction of the myofibrils. Histograms of fluctuations in naive rigor myofibrils examined
immediately after preparation were compared with histograms from myofibrils that had first contracted and then returned to rigor. The contribution of the second peak was

FIGURE 6 Examples of histograms of polarized fluorescence from contracting myofibrils. Horizontal scale: The negative value of fluorescence polarization. Vertical scale: The number of events during 20 s. The positions of minor (red line) and major (green line) peaks from a double Gaussian distribution fit
are (clockwise from the top left) P ¼ 0.3 and 0.11, P ¼ 0.29 and 0.06, P ¼ 0.2 and 0.06, P ¼ 0.33 and 0.14, P ¼ 0.32 and 0.11, and P ¼ 0.31
and .012. The ratios of the major peak (green line) to the corresponding minor peak (red line) in the double Gaussian fits are (clockwise, from the top left
corner) 2.7, 1.1, 0.5, 2.7, 1.6, and 3.1. The blue line is the sum of red and green line fits.
Biophysical Journal 100(4) 1024–1033

Pre- and Post-Power Stroke

1031

TABLE 2 Average polarization values from the Gaussian
distribution fits
Myofibrils
Rigor
EDC-rigor
Relaxation
EDC-relaxation
Contraction

Peak 1

Peak 2N

0.04 5 0.02
0.05 5 0.08
0.03 5 0.03
0.06 5 0.02
0.14 5 0.04

0.31 5 0.05

18
20
18
18
16*

Errors are given as SD, and N is the number of experiments.
*A total of 20 histograms were analyzed, and four of these were best fit bya single Gaussian distribution with the peak position corresponding to thatreported by peak 1.

small in both rigor cases (2.6 and 8.8%, respectively;
Fig. S5) in comparison with contracting myofibrils (51%;Fig. S6).
DISCUSSION
The main finding of this study is that polarization histograms from fluorescently labeled LC1 on the myosin head
during muscle contraction consist of two separate peaks.
Because the peaks correspond to the number of timesa cross-bridge assumes a given orientation during 20 s ofcontraction, it makes sense to suggest that during contraction, cross-bridges assume two distinct orientations. There
was a large difference in polarization between the two
peaks. They most likely indicate cross-bridge conformationsin the pre-and post-power-stroke positions. The first peakcorresponds to the pre-power-stroke position; the cleft
between the upper and lower subdomains of the 50 KDa
fragment is partially closed, the lever arm is in the up orientation and the myosin active site contains ADP and Pi (5).
The second peak corresponds to the post-power-stroke position when the cleft between the upper and lower subdomainsof the 50 KDa fragment is totally closed, the lever arm is in
the partially down orientation and the myosin active site
contains only ADP (5). The area corresponds to the average
number of times a cross-bridge assumes a given orientation
during the 20 s contraction, and the average ratio of the area
under peak 1 divided by the area under peak 2 was 5.7.
Therefore, the fact that the pre-power-stroke state is more
populated than the post-power-stroke state is consistent
with the notion that the transition from the pre- to thepost-power stroke is rate-limiting.
In a previous study, Ostap et al. (9) used EPR spectroscopy to detect ATP-induced changes in the structure ofspin-labeled myosin heads in whole muscle fibers in rigor
upon transition to relaxation and contraction. Different
conformational states were resolved on the basis of nanosecond rotational motion within the protein. In rigor, onlya single conformation was detected, in agreement with thefindings presented here. In relaxation and isometric contraction, the EPR spectrum showed two conformations anda small fraction of heads in the oriented, actin-bound conformation. Ostap et al.’s rigor results are consistent with thosereported here; however, the relaxation and contraction
results differ, most likely because they used whole muscle
fibers labeled at Cys-707, whereas we observed threecross-bridges labeled at LC1 in myofibrils. Ling et al. (12)and later Hopkins et al. (25) studied the breadth of the orientation distribution of fluorescent probes bound to the regulatory light chain of myosin, and found that it was essentiallythe same in different physiological states. This result is
consistent with our study, which also did not show consistent changes in the width of distributions.
In our experiments, the post-power-stroke conformationof the lever arm is different from the rigor conformation.
This implies that each cross-bridge executes a rotation in
two steps. Fig. 7 illustrates schematically the presumed
FIGURE 7 Schematic illustration of the crossbridge power stroke. In the pre-power-stroke state,corresponding to peak 1 of the polarized fluorescence histograms from contracting myofibrils, theupper and lower 50 K domains (UP50 and L50;
notation as in Coureux et al. (5)) are in a partiallyclosed conformation and LC1 is pointing sidewise(low polarization). In the post-power-stroke state,corresponding to peak 2 of the polarized fluorescence histograms from contracting myofibrils, theupper and lower 50 K domains are in a closed
conformation and LC1 is pointing downward(high polarization). The rigor state is short-lived
and the myosin head adopts a conformation similar
to that of the post-power-stroke state (5) exceptthat LC1 is now pointing completely sideways(zero polarization). This schematic does not take
into account such complicating factors as disorganization of actin filaments and inhomogeneities
between myofibrils. For simplicity, it is assumed
that the transition dipole of rhodamine is parallelto the long axis of LC1.
Biophysical Journal 100(4) 1024–1033

1032

sequence of events. A cross-bridge is initially in the prepower-stroke state, represented by peak 1, and subsequently
transits to the post-power-stroke state (weak ADP binding
state), represented by peak 2. If the rigor state (strong
interaction of the apo-head with actin) is a part of thecross-bridge cycle, there must be a second transition fromthe post-power-stroke state to the rigor state. The rigor
conformation is not seen during contraction because thisstate is short-lived in the presence of excess ATP. This
interpretation is consistent with the model originallyproposed by Huxley and Simmons (33), which suggeststhat the myosin head has a small number of sites, each ofwhich is capable of combining reversibly with a corresponding site on F-actin.
We did not attempt to translate the polarization values tothe absolute orientation of transition dipoles with respect tothe myofibrillar axis, because this orientation is criticallydependent on the model used to arrange the cross-bridges.
For example, if it is assumed that the cross-bridges arearranged helically along the long axis of muscle and thefluorescence contains a% contribution from a random, immobilized component, the difference between the pre- andpost-power-stroke angles will correspond to an angularchange from 90 to 5 depending on the value of a (22).
However, by calculating the average area under theGaussian curves, we can estimate that the relative populations of cross-bridges are ~70% in the pre-power-stroke
states and ~30% in the post-power-stroke states.
An additional possibility is that the two different peaks
indicate two populations of cross-bridges, each of which
executes a distinct rotation. This is unlikely, however,
because the presence of two populations was not detectedin direct electron microscopy measurements of cryofixed,
isometrically active, insect flight muscle (13). It is also
possible that cross-bridges do not rotate at all duringisometric contraction, and our results reflect their static
arrangement. In the absence of filament sliding, crossbridges may not be able to rotate while attached, andcross-linking may prevent them from reaching neighboring
actin target zones while detached (13). This interpretation
does not alter the conclusions of this study. The observed
distributions would then correspond to static positions thatwere assumed after the first molecule of ATP was split
when a cross-bridge was still allowed to rotate against an
unstretched elastic element.
We want to emphasize that, in contrast to previous studies(e.g., Muthu et al. (26)), we did not perform a correlation
analysis of the kinetic data. This is a significant advantage
of our method because a correlation analysis requires
assumptions about a model of cross-bridge-actin interaction.
We considered three possible artifacts, as described
below, and concluded that all are unlikely:
1. The second peak is due to rotation of the rhodamine
moiety alone. Although the mobile fraction was 10%
Biophysical Journal 100(4) 1024–1033

Midde et al.

(Fig. 3 B), this is unlikely because 1), rotational motion
on a scale of nanoseconds does not affect polarizationof fluorescence measured on a scale of milliseconds;
and 2), the nanosecond rotation of rhodamine is clearlyvisible in a solution of free dye (Fig. 3), whereas thesecond peak is not visible in rigor myofibrils (Fig. 5).
2. Addition of ATP alone is sufficient to induce the appearance of the peaks. However, in control experiments we
saw no effect of ATP on the anisotropy or fluorescent
lifetime of rhodamine-LC1.
3. The fluctuations are caused by myofibril movement, notcross-bridge rotations. This is unlikely because thecontrol experiments, in which we measured sarcomere
length as a function of the concentration of cross-linker,
showed no shortening upon addition of ATP when myofibrils were cross-linked with 20 mM EDC.

SUPPORTING MATERIAL
Six figures are available at http://www.biophysj.org/biophysj/supplemental/
S0006-3495(11)00052-X.
This work was supported by grants from the National Institutes of Health(R01AR048622 and R01HL090786).

REFERENCES
1. Geeves, M. A., and K. C. Holmes. 2005. The molecular mechanism ofmuscle contraction. Adv. Protein Chem. 71:161–193.
2. Houdusse, A., and H. L. Sweeney. 2001. Myosin motors: missing structures and hidden springs. Curr. Opin. Struct. Biol. 11:182–194.
3. Ferenczi, M. A., S. Y. Bershitsky, ., A. K. Tsaturyan. 2005. The ‘‘roll andlock’’ mechanism of force generation in muscle. Structure. 13:131–141.
4. De La Cruz, E. M., and E. M. Ostap. 2009. Kinetic and equilibrium
analysis of the myosin ATPase. Methods Enzymol. 455:157–192.
5. Coureux, P. D., H. L. Sweeney, and A. Houdusse. 2004. Three myosin
V structures delineate essential features of chemo-mechanical transduction. EMBO J. 23:4527–4537.
6. Sakamoto, T., M. R. Webb, ., J. R. Sellers. 2008. Direct observationof the mechanochemical coupling in myosin Va during processive
movement. Nature. 455:128–132.
7. Sellers, J. R., and C. Veigel. 2010. Direct observation of the myosin-Va
power stroke and its reversal. Nat. Struct. Mol. Biol. 17:590–595.
8. Walker, M. L., S. A. Burgess, ., P. J. Knight. 2000. Two-headed
binding of a processive myosin to F-actin. Nature. 405:804–807.
9. Ostap, E. M., V. A. Barnett, and D. D. Thomas. 1995. Resolution of
three structural states of spin-labeled myosin in contracting muscle.
Biophys. J. 69:177–188.
10. Thomas, D. D., and R. Cooke. 1980. Orientation of spin-labeled
myosin heads in glycerinated muscle fibers. Biophys. J. 32:891–906.
11. Borejdo, J., O. Assulin, ., S. Putnam. 1982. Cross-bridge orientationin skeletal muscle measured by linear dichroism of an extrinsic
chromophore. J. Mol. Biol. 158:391–414.
12. Ling, N., C. Shrimpton, ., M. Irving. 1996. Fluorescent probes of theorientation of myosin regulatory light chains in relaxed, rigor, andcontracting muscle. Biophys. J. 70:1836–1846.
13. Tregear, R. T., M. C. Reedy, ., M. K. Reedy. 2004. Cross-bridge
number, position, and angle in target zones of cryofixed isometrically
active insect flight muscle. Biophys. J. 86:3009–3019.

Pre- and Post-Power Stroke1033

14. Sabido-David, C., B. Brandmeier, ., M. Irving. 1998. Steady-state
fluorescence polarization studies of the orientation of myosin regulatory light chains in single skeletal muscle fibers using pure isomersof iodoacetamidotetramethylrhodamine. Biophys. J. 74:3083–3092.

24. Sabido-David, C., S. C. Hopkins, ., M. Irving. 1998. Orientation
changes of fluorescent probes at five sites on the myosin regulatory
light chain during contraction of single skeletal muscle fibres. J. Mol.
Biol. 279:387–402.

15. Hopkins, S. C., C. Sabido-David, ., Y. E. Goldman. 1998. Fluorescence polarization transients from rhodamine isomers on the myosin
regulatory light chain in skeletal muscle fibers. Biophys. J. 74:3093–
3110.

25. Hopkins, S. C., C. Sabido-David, ., Y. E. Goldman. 2002. Orientation
changes of the myosin light chain domain during filament sliding in
active and rigor muscle. J. Mol. Biol. 318:1275–1291.

16. Kraft, T., T. Mattei, ., B. Brenner. 2002. Structural features of crossbridges in isometrically contracting skeletal muscle. Biophys. J.
82:2536–2547.
17. Burghardt, T. P., T. Ando, and J. Borejdo. 1983. Evidence for crossbridge order in contraction of glycerinated skeletal muscle. Proc.
Natl. Acad. Sci. USA. 80:7515–7519.
18. Qian, H., S. Saffarian, and E. L. Elson. 2002. Concentration fluctuations in a mesoscopic oscillating chemical reaction system. Proc.
Natl. Acad. Sci. USA. 99:10376–10381.
19. Dos Remedios, C. G., R. G. Millikan, and M. F. Morales. 1972. Polarization of tryptophan fluorescence from single striated muscle fibers. A
molecular probe of contractile state. J. Gen. Physiol. 59:103–120.
20. Dos Remedios, C. G., R. G. Yount, and M. F. Morales. 1972. Individualstates in the cycle of muscle contraction. Proc. Natl. Acad. Sci. USA.
69:2542–2546.
21. Nihei, T., R. A. Mendelson, and J. Botts. 1974. Use of fluorescencepolarization to observe changes in attitude of S-1 moieties in muscle
fibers. Biophys. J. 14:236–242.

26. Muthu, P., P. Mettikolla, ., J. Borejdo. 2010. Single molecule kinetics
in the familial hypertrophic cardiomyopathy D166V mutant mouseheart. J. Mol. Cell. Cardiol. 48:989–998.
27. Borejdo, J., D. Szczesna-Cordary, ., N. Calander. 2010. Familial
hypertrophic cardiomyopathy can be characterized by a specific patternof orientation fluctuations of actin molecules. Biochemistry. 49:5269–
5277.
28. Herrmann, C., C. Lionne, ., T. Barman. 1994. Correlation of ActoS1,
myofibrillar, and muscle fiber ATPases. Biochemistry. 33:4148–4154.
29. Tsaturyan, A. K., S. Y. Bershitsky, ., M. A. Ferenczi. 1999. Structuralchanges in the actin-myosin cross-bridges associated with force generation induced by temperature jump in permeabilized frog muscle fibers.
Biophys. J. 77:354–372.
30. Borejdo, J., P. Muthu, ., T. P. Burghardt. 2007. Rotation of actinmonomers during isometric contraction of skeletal muscle. J. Biomed.
Opt. 12:014013.

22. Tregear, R. T., and R. A. Mendelson. 1975. Polarization from a helix offluorophores and its relation to that obtained from muscle. Biophys. J.
15:455–467.

31. Magde, D., E. L. Elson, and W. W. Webb. 1974. Fluorescence correlation spectroscopy. II. An experimental realization. Biopolymers.
13:29–61.
32. Elson, E. L. 2007. Introduction to FCS. In Short Course on Cellular andMolecular Fluorescence, Vol. 2. Z. Gryczynski, editor. University ofNorth Texas, Fort Worth. 1–10.

23. Morales, M. F. 1984. Calculation of the polarized fluorescence froma labeled muscle fiber. Proc. Natl. Acad. Sci. USA. 81:145–149.

33. Huxley, A. F., and R. M. Simmons. 1971. Proposed mechanism of force
generation in striated muscle. Nature. 233:533–538.

Biophysical Journal 100(4) 1024–1033