The converter domain modulates kinetic properties of
Drosophila myosin
Kimberly Palmiter
Littlefield1,
Douglas M.
Swank1,
Becky M.
Sanchez1,
Aileen F.
Knowles2,
David M.
Warshaw3, and
Sanford I.
Bernstein1
Departments of 1 Biology and
2 Chemistry, Molecular Biology Institute and Heart
Institute, San Diego State University, San Diego, California
92182-4614; and 3 Department of Molecular Physiology and
Biophysics, University of Vermont, Burlington, Vermont 05405
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ABSTRACT |
Recently the converter domain, an
integral part of the "mechanical element" common to all molecular
motors, was proposed to modulate the kinetic properties of
Drosophila chimeric myosin isoforms. Here we investigated
the molecular basis of actin filament velocity
(Vactin) changes previously observed with the
chimeric EMB-IC and IFI-EC myosin proteins [the embryonic body wall
muscle (EMB) and indirect flight muscle isoforms (IFI) with genetic
substitution of the IFI and EMB converter domains, respectively]. In
the laser trap assay the IFI and IFI-EC myosins generate the same
unitary step displacement (IFI = 7.3 ± 1.0 nm, IFI-EC = 5.8 ± 0.9 nm; means ± SE). Thus converter-mediated
differences in the kinetics of strong actin-myosin binding, rather than
the mechanical capabilities of the protein, must account for the
observed Vactin values. Basal and
actin-activated ATPase assays and skinned fiber mechanical experiments
definitively support a role for the converter domain in modulating the
kinetic properties of the myosin protein. We propose that the converter
domain kinetically couples the Pi and ADP release steps
that occur during the cross-bridge cycle.
actin-activated adenosine 5'-triphosphatase activity; unitary step
displacement; skinned fiber preparations; cross-bridge cycle; chemomechanical coupling
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INTRODUCTION |
IN DROSOPHILA
MELANOGASTER, myosin II isoforms are generated by
alternative splicing of six exon sets in a single myosin heavy chain
(MHC) gene (8). For example, the skeletal myosin isoforms found in the fast indirect flight (IFI) and slow embryonic body wall
(EMB) muscles share no common alternative exons, although regions of
the protein encoded by constitutively spliced exons are identical
(10, 36). Alternative exon usage, resulting in
muscle-specific myosin isoform expression, is responsible for defining
the kinetic and mechanical properties of particular muscle groups
(28). The most striking example is that of the indirect flight muscle (IFM) that contains the IFI myosin isoform. This specialized group of muscle fibers oscillates at the resonant frequency
of the flight system, generating wing beat frequencies on the order of
220 Hz and enabling the insect to fly (for review, see Refs.
13, 19). Embryonic body wall muscle
(containing the EMB isoform), on the other hand, is a slow muscle used
for larval locomotion and is histolyzed in the later stages of
morphogenesis. Muscle fibers isolated from transgenic flies expressing
the IFI and EMB myosin isoforms in the IFM and subjected to sinusoidal length oscillation experiments show profound mechanical and kinetic differences that are also apparent in isolated myosin preparations (28, 29). These results suggest that IFI and EMB myosin
isoforms confer very different biophysical and biochemical properties
on the contractile characteristics of the IFM. Furthermore, because these isoforms differ only in the alternatively encoded regions, this
suggests that the functional properties of the myosin are defined by at
least one of these regions.
The converter domain, a specialized region within the enzymatic
globular head of the MHC (see Fig. 1,
A and B), is thought to be involved in coupling
the energy of ATP hydrolysis to the mechanical events of the power
stroke. In Drosophila, the converter domain is encoded by
exon 11 (8). Exon 11 is one of the six alternatively spliced exon sets that code for variable regions in
Drosophila myosin isoforms. Four of the variable regions are
in the enzymatic S1 head (for review, see Ref. 3). The
converter regions in the IFI and EMB myosin isoforms are encoded by the
exon 11e and 11c isovariants, respectively, and are markedly different,
with 23 of 39 nonconserved amino acid substitutions (Fig.
1C). Transgenic Drosophila were created by
genetically swapping single alternative exons, including exon 11, between the IFI and EMB isoform backbones (29). The
naturally occurring IFI and EMB myosin isoforms and the chimeric IFI-EC
(IFI isoform backbone with the EMB converter domain) and EMB-IC (EMB
isoform backbone with IFI converter domain) myosins provide us with a
practical, integrative way to elucidate answers to complex
structure-function questions and a powerful approach to ascribe a
specific role to the converter domain in defining myosin function.

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Fig. 1.
Myosin converter domain and Drosophila myosin isoform
amino acid sequences. A: chicken skeletal S1 fragment
(25) with the converter domain (residues 712-779;
Ref. 7) highlighted in red and green. Residues
724-764 (highlighted in green) of the chicken skeletal myosin
correspond to those encoded by exon 11 in the Drosophila
myosin isoforms. Residues 731-738 are not resolved in this
structure. B: enlarged view of the converter domain.
C: amino acid sequence of the indirect flight muscle (IFI)
and embryonic body wall (EMB) isoforms encoded by exons 11e and 11c,
respectively. * Nonconserved amino acid residues between the IFI and
EMB converters.
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Alternative expression of IFI and EMB isoform-specific converter
domains was recently shown to play a role in defining the functional
properties of myosin. In vitro motility studies showed a dramatic
influence on actin sliding velocity after genetic substitution of this
region in the IFI-EC and EMB-IC chimeras (29). Similarly, mechanical studies on the IFM fibers expressing the chimeric proteins showed a dramatic influence on power-generating ability. The results of
these studies led to the speculation that the converter domain modulates the kinetic properties of Drosophila myosin.
In the present study, we assessed the molecular basis for the apparent
changes associated with converter domain substitution in the IFI-EC and
EMB-IC chimeras. We determined the mechanical and enzymatic properties
of the naturally occurring IFI and EMB isoforms and the chimeric IFI-EC
and EMB-IC myosins with single-molecule, solution ATPase, and fiber
mechanical assays. Using the single-molecule laser trap assay, we found
that the previously observed changes in actin filament velocity
(Vactin) (29), determined with the different myosin isoforms and chimeras, are not due to changes in the
inherent mechanical capacity of the myosin molecule. This is definitive
evidence that a kinetic mechanism must account for the converter
domain-mediated changes in Vactin. In support of this, solution biochemical actin-activated ATPase assays (the first of
their kind performed with native Drosophila myosin isoforms and chimeras) and skinned fiber data suggest that the converter domain
modulates actomyosin kinetics, influencing the duration of several
states in the cross-bridge cycle. We propose that interactions between
the motor core (actin binding and ATP hydrolysis center) and the
converter domain kinetically couple biochemical state transitions over
the duration of the ATPase cycle.
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EXPERIMENTAL PROCEDURES |
Drosophila myosin isolation.
Myosin was isolated from the IFM (dorsolongitudinal fibers) of
120-150 wild-type (IFI isoform) or transgenic (flies expressing the EMB, IFI-EC, or EMB-IC myosin in the IFM) flies as previously described (28) with the addition of one protease inhibitor
cocktail tablet (Roche Biochemicals, New York, NY) per 10 ml of each
purification solution. The final pellet of purified myosin was
resuspended in 25-50 µl of myosin storage buffer (MSB; 0.5 M
KCl, 20 mM MOPS, pH 7.0, 2 mM MgCl2, and 10 mM DTT). Myosin
concentration was determined by spectrophotometry using extinction
coefficient E
(1 mg/ml at 280 nm) = 0.53 cm
1 (17).
Actin isolation.
Actin was isolated from acetone powder prepared from chicken pectoralis
muscle and quantified as previously described (23). The
final pellet of purified F-actin was resuspended in actin storage
buffer (0.1 M KCl, 4 mM imidazole, 2 mM MgCl2, 0.5 mM ATP,
pH 7.0, 1 mM Na azide, 1 mM DTT). F-actin was stored on ice at 4°C
and used within 1 mo of preparation.
Laser trap experiments.
Flow cell construction, solution composition, and the laser trap setup
(including trap stiffness, quadrant detector calibration, and data
analysis) were previously described in detail (1, 9, 21).
Unitary displacement events were detected at low trap stiffness
(~0.02 pN/nm) as changes in the bright field image position of one of
the trapped beads projected onto a quadrant photodiode detector.
Displacement signals were obtained in both the x and
y directions, where x was parallel to the axis of
the actin filament. Displacement (Bessel filtered at 2 kHz) signals were digitized at 4 kHz and analyzed with the mean-variance (MV) analysis technique. Application of this analysis technique, originally developed for use in measuring single-ion channel currents and kinetics
(24), and for the determination of single-myosin molecule mechanical parameters, i.e., unitary step displacement amplitude (d), has been described in detail elsewhere (1, 9,
21). Briefly, MV analysis involves passing a time window over
the displacement data point by point and calculating a position mean
and variance for all points in that window. The mean and variance data
are then compiled as a three-dimensional histogram: mean
(x-axis), variance (y-axis), and counts
(z-axis). For clarity, MV histograms are presented in two
dimensions [i.e., mean (nm) and variance (nm2)] with the
total counts at a given mean and variance color-mapped on the
z-axis. Typically, MV histograms have two apparent regions of high density (populations) that can be attributed to baseline and
unitary events (see Fig. 2B).
The event population is statistically fit with a Gaussian distribution
in the x-direction (mean) and a
2
distribution in the y-direction (variance). Resolving power
is obtained through MV analysis, as event populations are offset from
the baseline population by the reduction in variance that occurs during
a unitary event. This decrease in variance occurs as myosin attaches to
actin, resulting in a reduction of the inherent noise attributable to
Brownian motion.

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Fig. 2.
Unitary step displacement
(d) data. A: raw displacement data obtained with
IFI-EC (IFI isoform backbone with EMB converter domain) myosin. Arrows
denote single unitary events. B: mean-variance (MV)
histogram generated from the complete data set (33 s) partially
illustrated in A. Baseline and event populations are denoted
with B and e, respectively. For this particular MV histogram, the event
population was fit at 5.3 ± 0.09 nm. C: d
(mean ± SE) obtained with the IFI and IFI-EC myosin
isoforms.
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Ca2+-ATPase assays.
High-salt Ca2+-ATPase assays were performed as previously
described (28).
Actin-activated ATPase assays.
Time course assays at a saturating actin concentration (5 µM) were
performed before the actin-activated ATPase assays to determine the
linear temporal range of phosphate (Pi) generation. These experiments (data not shown) were performed to ensure that the actin-activated ATPase assays were not substrate limited. Steady-state actin-activated ATPase assays were performed as previously described (15, 30) with the following modifications:
Drosophila myosin (100 nM) was preincubated for 10 min with
chicken F-actin (0-7.5 µM) in (mM) 20 KCl, 20 imidazole, 0.1 CaCl2, 5 MgCl2, pH 6.0, and 10 DTT at 25°C.
The ATPase reaction was started with the addition of NaATP to a final
concentration of 2 mM and allowed to proceed for 5 (IFI and IFI-EC) or
10 (EMB and EMB-IC) min. Blank reaction tubes containing only actin
(0-7.5 mM) were assayed simultaneously. Pi generation
was determined by extracting 50 µl of the ATPase reaction volume (150 µl total) into 500 µl of a 0.039% malachite green-1.1% ammonium
molybdate-1 N HCl (Sigma, St. Louis, MO)-0.02% Sterox (Bacharach,
Pittsburg, PA) solution. After 1 min the color development reaction was
quenched by the addition of 50 µl of 34% sodium citrate. Optical
densities (OD) were read at 650 nm with a Beckman Coulter DU640B
spectrophotometer (Fullerton, CA). Total Pi generated was
calculated by subtracting the appropriate actin blank OD from the
actin-myosin reaction OD and using the parameters generated in the
linear regression analysis of a Pi standard curve.
Muscle isolation and skinned fiber preparation.
A bundle of IFMs was removed from a Drosophila half-thorax
as described previously (28). To avoid the negative
influence of a deteriorating ultrastructure in the EMB and EMB-IC
lines, we used fibers from flies younger than the age at which
myofibril deterioration starts (<2 h old) (29). Using
fibers from young flies did not affect kinetics, as shown by the
identical shapes of the viscous modulus from 2-day-old and 2-h-old IFI
control lines (see Fig. 5). Thus we can compare kinetics across all
four fiber types. However, the amplitude of the viscous modulus is reduced in young flies because of the smaller myofibrillar cross sectional area per muscle cross-section (26).
The fibers were separated, and a single fiber was split lengthwise to
improve diffusion of ATP into the fiber during mechanical experiments.
The preparations were ~100 µm in diameter and ~0.6 mm in length.
Fibers were chemically demembranated (skinned) in a relaxing solution
containing 5 mM MgATP, 15 mM creatine phosphate, 240 U/ml creatine
phosphokinase, 1 mM free Mg2+, 5 mM EGTA, 20 mM
N,N-bis(z-hydroethyl)-zaminoethane sulfonic acid
(BES) (pH 7.0), 1 mM DTT, and a protease inhibitor cocktail (Roche
Biochemicals) containing 0.5% Triton X-100 and 50% glycerol for
1 h at 4°C. The ionic strength was adjusted with sodium
methane sulfonate to 200 mM. Aluminum T clips were used to mount the
fibers on the mechanical rig.
Determination of rate of tension redevelopment.
To determine the rate of tension redevelopment, fibers were subjected
to a series of four identical 0.5% muscle-lengthening steps. The force
response was averaged over the four steps. The resulting phase
3 of the force response, r3
(20) [equivalent to rate of tension development after a
quick stretch (KTR) (5)], was fit
with an exponential rise to a maximum: y = yo + a (1
e
bx), where b is
r3.
Determination of frequency of maximal work per cycle.
To determine the frequency of maximal work per cycle
(Wfmax), fibers were subjected to sinusoidal length
oscillation experiments, the details of which were previously described
(6). Briefly, the fiber was activated stepwise by
progressively exchanging the initial relaxing solution with activating
(pCa 4.0) solution, up to pCa 4.5. Sinusoidal length changes of 0.25%
muscle length (full amplitude) were applied over 47 frequencies from 1 to 1,000 Hz. For each frequency, elastic and viscous moduli were
calculated from the force response to sinusoidal length perturbations
by computing the amplitude ratio and the phase difference for force and
length and dividing the ratio by fiber cross-sectional area. Temperature was 15°C for all mechanics experiments.
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RESULTS |
Single-molecule mechanics show that unitary step displacement
amplitudes of IFI and IFI-EC myosins are the same.
We determined the amplitude of the unitary step displacement,
d, generated by both the IFI and IFI-EC myosins with the
laser trap assay. Figure 2A shows 3 s of raw
displacement data obtained with IFI-EC myosin in 3 µM ATP. Arrows
denote single unitary events. The MV histogram (see EXPERIMENTAL
PROCEDURES for detailed description) generated from the complete
raw data set partially illustrated in Fig. 2A is shown in
Fig. 2B. Baseline (the time during which myosin is detached
from actin) and event (duration for which actin and myosin are strongly
associated) populations are denoted by B and e, respectively. Baseline
data have a higher position variance (owing to the effects of Brownian
motion) and an average displacement of 0 nm. Event populations, on the
other hand, are characterized by a lower variance due to stiffening of
the system when myosin attaches to actin. Average d values
(Fig. 2C) were 7.3 ± 1.0 (n = 9) and
5.8 ± 0.9 (n = 11) nm (means ± SE) for the
IFI and IFI-EC isoforms, respectively, and are not significantly
different by Student's t-test.
ATPase activity increased in both chimeras.
The enzymatic activity of the native myosin isoforms and chimeras was
determined both in the presence (actin-activated ATPase) (Fig.
3 and Table
1) and absence (basal
Ca2+- and Mg2+-activated ATPase; Table 1) of
actin. The actin-activated ATPase activity of the IFI myosin isoform
was significantly greater than the EMB isoform. This
difference in actin-activated ATPase activity is inherent to
the IFI and EMB myosins as reflected in both their basal
Ca2+- and Mg2+-activated ATPase activities
(Table 1). Interestingly, however, the Vmax of
EMB myosin was potentiated 10-fold by actin, whereas IFI activity was
potentiated only ~4-fold. There was no difference in the
Km between these myosins.

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Fig. 3.
Steady-state rates of actin-activated myosin ATPase.
A: IFI vs. IFI-EC. B: EMB vs. EMB-IC (EMB isoform
backbone with IFI converter domain). P < 0.05 by
Z-test on the parameter fits for IFI vs. IFI-EC ( ) and
EMB vs. EMB-IC (**).
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In contrast, the chimeric myosins had actin-activated ATPase activities
that were, in both cases, greater than their respective control myosins
(Fig. 3, A and B). We observed similar increases in both the basal Ca2+- and Mg2+-activated
ATPase activities for the IFI-EC relative to the IFI myosin. However,
basal rates for the EMB-IC chimera, relative to the EMB isoform, did
not increase. We observed approximately twofold decrease in the
Km for actin in the EMB-IC myosin as well as the
IFI-EC chimeric myosin (see Table 1 for all data).
Skinned fiber data support a kinetic role for the converter domain.
All fibers exhibited the classic delayed tension rise following a quick
stretch that promotes work generation in IFM (Fig. 4). However, fibers expressing EMB,
EMB-IC, and IFI-EC myosin vary dramatically in the rate of tension
redevelopment after stretch (r3) (Table 2 and
Ref. 29). The r3
values are 11.8-fold lower in fibers containing EMB myosin compared
with the IFI fibers. The r3 values were lower
for IFI-EC fibers relative to the IFI and higher in the EMB-IC fibers
compared with EMB.

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Fig. 4.
Rates of tension redevelopment (r3). Fully
activated indirect flight muscle (IFM) skinned fibers were subjected to
a rapid lengthening step [0.5% muscle length (ML)], after which the
rate of force redevelopment was measured. Open circles are
representative traces from the IFI, EMB, IFI-EC, and EMB-IC fiber
types. Solid lines are the fit of the data.
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Fibers were also subjected to sinusoidal length oscillation experiments
(Fig. 5). Because we are interested in
the kinetics of active force generation by myosin cross bridges, we
focused on the viscous modulus, which reveals the optimal frequency
range for work generation by cross bridges. The four fiber types differ dramatically in the frequency range over which useful work is generated
(by convention, where the viscous modulus is negative). For example, in
EMB fibers the modulus is negative from 2 to 25 Hz, in EMB-IC fibers
from 5 to 80 Hz, in IFI-EC fibers from 8 to 150 Hz, and in wild-type
fibers from 15 to 230 Hz. The lowest point of the viscous modulus curve
corresponds to the Wfmax. The Wfmax values
agree in relative order and magnitude with the
r3 measurements.

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Fig. 5.
Viscous moduli of maximally Ca2+-activated
IFM skinned fiber preparations vs. frequency. Graphs depict the
frequency ranges in which there is a phase lag of the tension with
respect to the length. A: viscous modulus of IFM skinned
fiber preparations obtained from Drosophila 2 days after
eclosion (IFI isoform; n = 6) and IFI-EC-containing
fiber preparations (n = 6). B: viscous
modulus of IFM skinned fiber preparations obtained from
Drosophila <2 h after eclosion (IFI isoform), fibers
containing the EMB (n = 7) or EMB-IC (n = 6) myosin. See Table 2 for r3 and frequency of
maximal work (Wfmax) data.
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DISCUSSION |
Our purified chimeric myosin and fiber preparations had the
converter region exchanged between two naturally occurring
Drosophila myosin isoforms (IFI and EMB). The relay helix,
converter domain, and light chain-binding domain (putative
lever arm) make up the proposed "mechanical element"
(32). A series of structural transitions within these
specialized domains results in the generation of the power stroke. The
ability to express chimeric myosin, where changes in primary sequence
and (possibly) structure are limited to a defined region of the
molecule, offers the opportunity to ascribe specific functional
properties to a particular structural domain. In this study we
provided direct evidence that the converter domain influences
myosin's kinetic properties by performing single-myosin molecule step
displacement, enzymatic ATPase, and skinned fiber length oscillation experiments.
Mechanical properties of IFI and IFI-EC myosin proteins.
We have demonstrated that the 2.4-fold attenuation of
Vactin previously observed with the IFI-EC
chimera (29) cannot be accounted for by a change in the
inherent mechanical capacity of the myosin molecule. At the molecular
level Vactin
d/ton (12) where
d is the unitary step displacement and
ton is the duration of the strong actin-myosin
interaction. Given that the amplitude of myosin's unitary step is
constant for three of the four isoforms characterized in this and a
previous study (28), the observed differences in
Vactin must be related to differences in the
time spent in the strongly bound state following the power stroke
(ton), as has been observed in other myosin
isoforms (16, 21, 22, 30). Thus swapping converter domains
does not perturb the mechanical capabilities of the
Drosophila myosin but rather the kinetics of steps in the
actomyosin ATPase cycle that contribute to the duration of
ton.
ton is determined by the rates of both ADP
release (k
ADP) from, and ATP binding
(k+ATP[ATP]) to, the myosin active site
(1, 15, 21, 31). For Drosophila myosin the ADP
release rate is most likely faster than that of chicken skeletal muscle myosin [~500 s
1 (18, 27)]. This would
correspond to an average ton of <2 ms at
saturating ATP concentrations in the laser trap assay. Given that this
duration is less than the temporal resolution of the assay, it would be
impossible to determine whether the ton values for the IFI and IFI-EC myosins differ at saturating ATP, thus preventing us from relating any potential differences in
ton to changes in the ADP release rate.
Solution kinetic measurements.
Characterizing the actin-activated ATPase activity of the IFI and EMB
isoforms and the two chimeras provides additional, novel insight into
the kinetic properties of the myosin isoforms and chimeras. The low
rate of ATPase activity observed specifically with the IFI isoform is
consistent with previous actin-activated ATPase rates (~2
s
1) determined in Drosophila skinned fiber
preparations (34). The expected differences in ATPase
activity observed with the IFI and EMB myosins may represent functional
tuning of the myosin specific for the muscle environment in which they
are expressed. The enhanced actin-activated ATPase activities of both
chimeric myosins relative to their controls may reflect the ability of actin to provide a greater level of activation relative to either the
EMB or IFI myosin. Alternatively, it is likely that genetic swapping of
the fast IFI and slow EMB converter domains perturbs normal structural
interactions that constrain the rate of ATP hydrolysis and presumably
Pi release. Finally, the increase in ATPase activity of the
IFI-EC chimera, above that of the fastest IFI isoform, was unexpected
but other instances of myosin mutations causing an increase in ATPase
have been reported (30, 35).
Duty ratio.
A kinetic parameter frequently used to characterize myosin is its duty
ratio (f), defined as the fraction of the total cycle time
(tcycle) that myosin remains strongly bound to
actin after the power stroke, where
Although ton and
tcycle vary across the vertebrate class II
muscle myosin isoforms, the ratio of ton to
tcycle, f, is constant (31). Both ton and
tcycle can be estimated from
Vactin and the ATPase rate, respectively, where
so that
Because d is constant across the various
Drosophila myosin isoforms, then
Using this relationship, we estimated a relative duty ratio for
the two isoforms and two chimeras (see Table
3). In contrast to the vertebrate muscle
myosins (31), our estimates suggest that the duty ratio is
not constant across Drosophila isoforms and chimeras. Given
that these estimates are model dependent, it is still possible that
differences between the IFI and EMB isoforms may have evolved so that
actomyosin kinetics are adapted to match the functional demands on the
myosin isoform. In both chimeras, the duty ratio estimates suggest that
the kinetic parameters that define f are uncoupled.
Interestingly, exchanging converter domains resulted in both chimeras
having a duty ratio closer in value to their respective donor
isoform (Table 3).
Average force generation.
Changes in f may have important implications for average
force generation (F) in skinned fiber preparations as
where N is the number of cross bridges and
Funi is the cross-bridge unitary force (33).
If we assume that N and Funi are constant for
the various fibers, then the average fiber force will be directly
proportional to the myosin duty ratio. This prediction is surprisingly
accurate with the rank order of duty ratios following that of the
maximum isometric force measured in skinned fiber preparations, i.e.,
FEMB > FIFI-EC > FIFI
(29). However, caution is warranted when making
predictions concerning isometric parameters based on measurements in
the motility assay or in solution where the actomyosin interaction
occurs under unloaded conditions.
Kinetic correlations.
When particular experimental parameters measured for myosin and muscle
fibers are coupled, or otherwise related (limited by the same
biochemical rate constants), they should correlate. For example, Tyska
and Warshaw (31) show a strong linear correlation between
Vactin and ATPase activity in the vertebrate
class II muscle myosins as was originally described in whole muscle
(2). Although the kinetic steps that govern
Vactin and ATPase activity are different (i.e.,
cross-bridge detachment rate for Vactin vs. attachment rate
for ATPase), a strong correlation between Vactin and the ATPase rate for the various vertebrate myosins argues that
changes in the kinetics of these steps are coupled as a result of
evolutionary selection. As illustrated in Fig.
6A, this does not appear to be
the case for the Drosophila myosin isoforms and chimeras.
The lack of correlation between Vactin and
ATPase activity (R2 = 0.009) suggests that
the converter can modulate the kinetic properties of multiple
rate-limiting steps in the actomyosin ATPase cycle to various extents.
However, conclusions about coupling in the native isoforms are
premature until additional native isoforms have been characterized.

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Fig. 6.
Kinetic parameter correlations. Previous actin filament velocity
(Vactin) data (29) were normalized
to those obtained with the IFI isoform. A:
Vactin vs. relative ATPase activity,
R2 = 0.009. B:
Wfmax vs. r3,
R2 = 0.87. C:
r3 vs. relative ATPase activity,
R2 = 0.26. D:
r3 vs. relative Vactin,
R2 = 0.47. Regression lines were generated
with SigmaPlot version 4.0 (SPSS, Chicago, IL).
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To understand how muscle fiber kinetics are influenced by the kinetic
changes seen at the molecular level, we correlated the rate of tension
recovery (r3) and the frequency for maximum work production (Wfmax) in skinned fibers to myosin ATPase
activity and Vactin. A strong linear correlation
(R2 = 0.87) between Wfmax and
r3 (Fig. 6B) is expected because
these parameters are governed by similar cross-bridge kinetic rate
constants [r3
2
(Wfmax)]
(14). Using the classic two-state model (11), r3 has been proposed to be influenced by both
cross-bridge attachment (f) and detachment (g) rates
(r3
f + g; Ref. 4). When
correlating fiber and molecular level parameters, we see no correlation
(R2 = 0.26) between
r3 and ATPase (Fig. 6C) activity,
although a weak correlation (R2 = 0.47)
does exist between r3 and
Vactin (Fig. 6D). This suggests that
although both attachment and detachment kinetics determine r3 (4), r3
is more strongly influenced by the same kinetic step that governs
Vactin, i.e., the rate of ADP release or
cross-bridge detachment. Again, although dramatic changes in both
molecular and fiber level properties can be achieved by exchanging the
converter domain, correlations between isolated myosin and fiber
kinetics should be made with caution given the influence of strain and a constraining lattice on fiber kinetics.
In conclusion, we have shown, by determination of the single-molecule
mechanical properties and enzymatic ATPase activity of native and
chimeric Drosophila myosin proteins, that the converter domain influences myosin's kinetic properties rather than its mechanical capabilities. These kinetic changes translate to the fiber level, resulting in a significant alteration in the kinetic properties of skinned fiber preparations. However, the structural mechanism by which changes to the converter domain modulate myosin's kinetic properties remains to be elucidated. It is apparent from both
the isolated myosin and fiber studies that the converter domain is not
the sole determinant of Drosophila myosin kinetics, because
swapping converter domains does not fully switch the myosin kinetics of
either chimera to that of the IFI or EMB isoform. Because there are
three other variable domains located in the myosin S1 head that could
influence myosin kinetics (for review see Ref. 3),
additional chimeras with single or combinations of alternative exons
will help identify which are required for interconversion between the
IFI and EMB isoforms.
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ACKNOWLEDGEMENTS |
We acknowledge the excellent technical assistance of Jennifer
Suggs, Brandon Walsh, Allen Church, and Massoud Nikkhoy. We thank Drs.
Ryan Littlefield, Mark Miller, Jeff Moore, and Brad Palmer for helpful
scientific discussions. In addition, we gratefully acknowledge Dr.
David W. Maughan for providing support and facilities for the fiber
mechanics experiments.
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FOOTNOTES |
This work was supported by National Institutes of Health Grants
GM-32443 (to S. I. Bernstein), HL-66157 (to D. M. Warshaw), and HL-68034 (to D. W. Maughan), and American Heart Association Western States Affiliate Fellowships 0120127Y (to K. P. Littlefield) and 0120022Y (to D. M. Swank).
Address for reprint requests and other correspondence:
K. P. Littlefield, Scripps Research Inst., Dept. of Cell
Biology, 10550 Torrey Pines Rd., La Jolla, CA 92037 (E-mail:
kplittle{at}scripps.edu).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published December 11, 2002;10.1152/ajpcell.00474.2002
Received 10 October 2002; accepted in final form 6 December 2002.
 |
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