Flight muscle properties and aerodynamic performance of Drosophila expressing a flightin transgene
1 Department of Biology, University of Vermont, Burlington, VT 05405,
USA
2 Department of Molecular Physiology and Biophysics, University of Vermont,
Burlington, VT 05405, USA
3 Biofuture Research Group, Department of Neurobiology, University of Ulm,
Albert-Einstein-Allee 11, 89081 Ulm, Germany
* Author for correspondence (e-mail: jim.vigoreaux{at}uvm.edu)
Accepted 1 December 2004
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Summary |
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Key words: insect flight muscle, flightin, thick filaments, stretch activation
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Introduction |
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Electron microscopy studies have provided insight into the precise and
ordered manner by which the myofilament lattice of Drosophila IFM is
assembled throughout development (Reedy
and Beall, 1993; Vigoreaux and
Swank, 2004
). Genetic approaches have been instrumental in
elucidating the role of myofibrillar proteins on sarcomere assembly and muscle
structure stability (for reviews see:
Bernstein et al., 1993
;
Cripps, 2004
;
Vigoreaux, 2001
). In
particular, Mhc gene mutants have provided insight into the role of
MHC protein domains in flight muscle development and function (for reviews
see: Miller and Bernstein,
2004
; Swank et al.,
2000
). Analysis of flightin gene mutants also have shown
that flightin plays an essential role in thick filament formation and flight
muscle function (Reedy et al.,
2000
; Vigoreaux et al.,
1998
).
Drosophila flightin is a novel 20 kDa IFM-specific protein, that
undergoes extensive phosphorylation during the late pupal stages of
development and throughout the initial hours of adulthood, preceding the
acquisition of flight (Vigoreaux and
Perry, 1994; Vigoreaux et al.,
1993
). It has been shown that flightin is a component of the thick
filament that, in vitro, binds the myosin rod
(Ayer and Vigoreaux, 2003
). A
single amino acid substitution in the myosin rod (Glu 1554 to Lys, the
Mhc13 allele) prevents the accumulation of flightin in
vivo and its binding to MHC in vitro
(Ayer and Vigoreaux, 2003
;
Kronert et al., 1995
).
Mhc13 flies are flightless and their IFM undergoes a
time-dependent hypercontraction that is characterized by myosin proteolysis,
thick filament instability and sarcomere degeneration
(Kronert et al., 1995
). A more
recent study showed that a null mutation in the flightin
(fln) gene, fln0, leads to a remarkably similar
phenotype as Mhc13 suggesting that the absence of flightin
severely compromises IFM structure and function
(Reedy et al., 2000
). In
addition, sarcomeres and thick filaments are longer than normal in IFM
suggesting that flightin plays a key role in thick filament length
determination. Mechanical analysis of skinned fibers from newly eclosed
fln0 and Mhc13 flies showed similar
deficits in passive and dynamic stiffness, and a loss of the stretch
activation response that resulted in no net positive work output
(Henkin et al., 2004
).
Together with studies that showed flightin is distributed throughout the
A-band of the sarcomere, these results suggest that flightin influences the
viscoelastic properties of the thick filaments. To test the hypothesis that
the ultrastructural and functional defects in fln0 are
attributed to the absence of flightin, we conducted the present study using
genetic transformation of wild-type
(fln+/fln+) and flightin null
(fln0/fln0) Drosophila using
a chimeric Actin88F-promoterfln gene construct. We
show that the transgene successfully rescued the ultrastructural and
contractile defects engendered by fln0 but the transgenic
flies do not recover full flight competency. We also found that increasing the
number of flightin genes to four has no adverse effect on IFM
properties.
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Materials and methods |
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Construction of transformation vector
To construct a transformation clone containing the flightin gene,
we started with pW8-Act88F-3'Tm2, a P-element transformation
vector derived from pW8/Tm2-35 (Miller et
al., 1993) and obtained from Terese Tansey. This vector contains
the Actin88F promoter region (extending from the XbaI site
at 1420 to the G just 5' of the initiation codon
(Geyer and Fyrberg, 1986
;
Klemenz et al., 1987
;
Rubin and Spradling, 1983
;
Sanchez et al., 1983
), part of
the multiple cloning site from pW8, and Tropomyosin (Tm2)
sequence from the 3' untranslated region. Using KpnI and
PstI restriction enzymes, the 3'-end of the
Tropomyosin gene was excised from the vector and replaced with a 1.14
kb KpnIPstI flightin genomic fragment
obtained from a
phage genomic library. This fragment extends from the
flightin start codon to 0.55 kb past the translation stop codon. The
phage library clone does not contain either the first untranslated
exon or intron 1. This Actin88Ffln chimeric gene was excised
from pW8 using EcoRI and PstI restriction enzymes and
subcloned into the P-element mediated transformation vector pCaSpeR
(Flybase #FBmc0000168).
Generation of transgenic lines
Transformation was performed as described elsewhere
(Spradling and Rubin, 1982).
The helper plasmid used was pUChs
23 (Flybase #0000938), which
was obtained from Margarita Cervera. The pCasPeR plasmid vector and helper
plasmid were amplified in XL1 Blue E. coli cells (Novagen, Madison,
WI, USA) in LB broth and purified using Qiagen maxiprep kit (Qiagen, Valencia,
CA, USA). The concentration of DNA injected was 0.39 mg ml-1 of
pCasPeR and 0.171 mg ml-1 of
23. Transformants were
identified by yellow or orange eye color in the G1 generation. Homozygous
strains were produced from crosses of individuals with darker eye color that
resulted in no white eye progeny. Each transgene was mapped to its
resident chromosome using w*; T(2;3)apXa,
apXa/CyO; TM3, Sb1 by standard crossing techniques.
The transgene was crossed into the fln0 strain using
standard crossing techniques.
Gel electrophoresis and western blot analysis
Denaturing one-dimensional (1DE) gel electrophoresis was performed using
the discontinuous buffer system (Laemmli,
1970) as described previously
(Vigoreaux et al., 1991
).
Two-dimensional (2DE) gel electrophoresis was performed using the Protean IEF
cell (BioRad Inc., Hercules, CA, USA). IEF strips (pH 47 gradient) were
used for the first dimension and precast 12.5% gels were used for the second
dimension. Separation in the first dimension was carried out using a three
step protocol. The IEF strips were rehydrated for 12 h at 20°C. Step two
involved a 2 h rapid volt ramp to 3500 V h-1 and step three focused
the strips for 14 h or 50,000 volt hours. To prepare samples for
electrophoresis, flies were placed in acetone for 1 h at room temperature
followed by lyophilization in a speed vac. The thorax was dissected away from
other body parts and homogenized in IEF sample buffer and spun down to remove
the cuticle debris.
Western blots were performed as described in Vigoreaux et al.
(1993) with an anti-flightin
polyclonal antibody described in Reedy et al.
(2000
). For developmental
blots, pupae were staged according to Bainbridge and Bownes
(1981
) and homogenized in
Laemmli sample buffer with 8 mol l-1 urea and protease inhibitors
(Vigoreaux et al., 1991
).
Samples were run on a 12% SDS gel, blotted onto membrane and processed for
antibody detection as described (Vigoreaux
et al., 1993
).
Protein expression assays
To determine relative expression levels, whole thoraces were homogenized as
described above. Protein concentration was determined using the BioRad DC
protein quantification kit and equal amounts of protein were loaded in
individual lanes of a 12% SDS gel. After electrophoresis, proteins were
transferred to Bio-Rad PVDF membrane and the membranes were blocked with Aqua
Block (East Coast Biologics, Inc., North Berwick, ME, USA). The primary
antibody was an anti-flightin polyclonal
(Reedy et al., 2000) and
Alexaflor 698 fluorescent antibodies (Molecular Probes, Eugene, OR, USA) were
used as secondary antibodies. After staining and washing, the membranes were
scanned on an Odyssey fluorescent scanner (LI-COR Biosciences, Lincoln, NE,
USA) and analyzed with Phoretix 1D software (Nonlinear Dynamics, Durham, NC,
USA) as follows. Each image was first converted from color to grayscale in
Photoshop and opened as a new experiment. After automatic selection of lanes,
the bands were manually selected and their borders adjusted based on the peak
profile in the analysis window. Protein quantity was obtained from band volume
after background subtraction.
Transmission electron microscopy
Fly thoraces were bisected and the separated halves were fixed for 2 h in
2.5% glutaraldehyde and 0.1% paraformaldehyde. After fixation samples were
stored in 0.1 mol l-1 Millonigs phosphate buffer, pH 7.2. Samples
were dehydrated through a series of ethanol from 35% through absolute for 10
min in each concentration. The final dehydration step was in propylene oxide 3
x for 5 min each. Infiltration was performed with propylene oxide and
Spurr's resin 3:1 for 30 min, 1:1 for 30 min, 1:3 for 45 min and 100% Spurr's
resin for 45 min. Embedding was done in 100% Spurr's resin and polymerized for
24 h at 70°C. Semi-thin sections (1 µm) were cut with glass knives on a
Reichert ultracut microtome, stained with methylene blue (azure II), and
evaluated for areas of interest. Ultrathin sections (6080 nm) were cut
with a diamond knife, retrieved onto 150 mesh copper grids, contrasted with
uranyl acetate (2% in 50% ethanol) and lead citrate, and examined with a JEOL
1210 TEM (JEOL USA, Inc., Peabody, MA, USA) operating at 60 kV.
Polarized light microscopy
Examination of IFM fiber morphology was done essentially as described
previously (Nongthomba and Ramachandra,
1999). Whole flies were dehydrated through a series of 50, 70, 80,
90 and 100% ethanol for 1 h in each solution at room temperature. The flies
were then placed in methyl salicylate for one hour at room temperature, fixed
on slides using permount and viewed under polarized light. Pictures were taken
with a digital camera and Magnafire imaging software (Oreko, Dulles, VA,
USA).
Flight test and wing-beat-frequency analysis
Flight test analysis and wing-beat-frequency analysis were performed as
described previously (Vigoreaux et al.,
1998).
Sinusoidal analysis of skinned flight muscle fibers
The sinusoidal procedure was performed as described previously
(Dickinson et al., 1997) using
IFM fibers from the dorsolongitudinal muscle (DLM), with the exception that
after being stretched to just taught, each fiber was stretched by 2%
increments until the oscillatory work (B component) was maximized, defined as
a <3% increase in B. The solutions used are described in Henkin et al.
(2004
).
In vivo estimates of kinematic and muscle performance
To evaluate muscle mechanical power output in an intact fly in flight, we
used an improved version of a method described elsewhere
(Lehmann and Dickinson, 1998;
Lehmann and Dickinson, 1997
).
Here we present only a brief description of the experimental procedure and
focus mainly on the differences from the previous studies. Female fruit flies
were tethered and flown in a virtual reality flight arena in which stroke
amplitude, stroke frequency, total force production and carbon dioxide release
were measured simultaneously. The flies actively modulated the azimuth
velocity of a vertical dark stripe displayed in the arena using the relative
difference in stroke amplitude between the two beating wings (closed-loop
feedback conditions). While flying in closed-loop, the animals typically
modulate kinematic and respirometric parameters in response to the motion of
an open-loop stripe grating (horizontal stripes) that were oscillated
vertically around the fly with a sinusoidal velocity profile. We have shown
previously that under those conditions, fruit flies may maximize their
locomotor output allowing the evaluation of maximum locomotor capacity
(Lehmann and Dickinson, 1997
).
We employed flow-through respirometry with a flow rate of 1000 ml
min-1 and used a Li-cor 7000 gas analyzer to measure the rate of
carbon dioxide release during flight. Compared with previous studies on flight
energetics in Drosophila, the higher flow rate yielded better
temporal resolution of the metabolic measures permitting a tighter correlation
between flight force and carbon dioxide production measures. We estimated the
temporal shift of the CO2 signal, due to the delay of the
connecting gas tubings and the washout characteristics of the respirometric
chamber, by performing cross-correlation between the force and CO2
signal. Since a previous study on flight energetics in Drosophila has
shown a transient effect on flight parameters following take-off, we excluded
the initial 5 s and the last 2 s of flight time within each flight sequence
(Lehmann and Dickinson, 1997
).
The average flight time of the three tested fly lines (wild-type Canton S,
P{fln+}fln0,
P{fln+}fln+) was 1747±680,
1271±93 and 1247±83 s (mean ± S.D.,
N=11, 15, 11), respectively. The ambient temperature was similar in
all experiments, approximately 23.9±1.0°C (t-test,
P>0.05).
To derive muscle efficiency in the flying animal, we estimated the power
requirements for flapping flight according to a set of equations and
parameters published previously (Lehmann
and Dickinson, 1998; Lehmann
and Dickinson, 1997
). Assuming 100% elastic energy storage within
an entire flapping cycle, muscle efficiency is given by the ratio between the
sum of induced power and profile power requirements for flight, and metabolic
power due to ATP conversion. It is difficult to derive exact values for
profile power because this measure critically depends on the drag coefficient
of the flapping wings that varies with wing kinematics. Previous studies
derived drag coefficient from Reynolds number assuming that a decrease in wing
flapping velocity is correlated with an increase in drag coefficient
(Lehmann and Dickinson, 1997
).
By contrast, here we estimated drag coefficient from lift coefficient values
employed by the tethered animal during flight. The latter coefficient can be
calculated from wing velocity and force measurements using conventional
aerodynamic theory (Ellington,
1984
; Lehmann and Dickinson,
1998
). Subsequently, the drag coefficient in the flying animal was
derived from lift (CL) and drag coefficient
(CD) polars measured in a dynamically scaled 3D
Drosophila robotic wing using the following equations:
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Results |
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We generated 11 independent transgenic lines, two on the X chromosome, two on the second chromosome, five on the third chromosome, and an additional two that have not been mapped. Three of these strains were crossed to fln0 and tested for flight. We focused on the strain with the best flight score, a transgenic line with an X chromosome insertion, P{Act88F-fln+}; fln0e-2 (hereby referred to as P{fln+}0.2). Table 1 summarizes the results of flight test of normal and P{fln+}0.2 transgenic flies. Note the significant improvement in flight performance of the rescued strain vs fln0. However, flight ability is not fully restored.
|
A characteristic feature of fln0 IFM is that fibers
hypercontract, resulting in detachment of one or both ends of the fiber from
the cuticle and `bunching' of the muscle mass
(Reedy et al., 2000). We
inspected IFM fibers of the P{fln+}0.2 rescued
lines by polarized light microscopy and determined their morphology to be
normal (not shown). A more-detailed analysis of sarcomere structure was
conducted by electron microscopy. On longitudinal sections sarcomeres appear
normal, with well-defined Z-bands and clearly depicted A-bands and I-bands
(Fig. 2). Sarcomere length is
uniform and similar to sarcomere length in wild-type flies
(Table 1 and
Fig. 2). On cross sections,
myofibrils from P{fln+}0.2 rescued flies are
circular with well-defined diameter, and show the normal double hexagonal
array of interdigitated thick filaments and thin filaments
(Fig. 2). However, there is a
decreased number of thick filaments per myofibril
(Table 1). Wild-type myofibrils
had an average of 945 thick filaments per myofibril, while
P{fln+}0.2 had significantly fewer with 782, a
decrease of about 17%. The number of myofibrils per area was no different in
the transgenic vs the control (not shown).
|
While the majority of the myofibrils appear normal, we did observe occasional myofibril defects not commonly seen in IFM of wild-type flies (Fig. 2I and 2J). Broken down sarcomeres with partially torn Z-bands and missing M-lines are seen, as well as myofibrils with fractures that suggest a faultily assembled lattice.
The reduced number of thick filaments and the reduced flight ability could
result from incorrect timing of gene expression, incorrect timing of
phosphorylation, and/or insufficient flightin expression. We tested
all three of these possibilities by conducting western blot analysis of
flightin accumulation and phosphorylation throughout pupal development and in
adults. We first looked at the developmental expression of the
flightin transgene. In wild-type flies, flightin begins to accumulate
at pupal stage P8 (60 h after pupation and
22 h after initial
myofibrils appear at 25°C; Vigoreaux
and Swank, 2004
) and continues to accumulate at increasing levels
into adulthood (Vigoreaux et al.,
1993
). An almost identical profile is seen in
P{fln+}0.2 transgenic flies except that
expression starts at the P7 stage, or 28 h before expression of the
endogenous protein normally begins (Fig.
3). This earlier onset of expression is not unexpected given that
the Act88F promoter is activated during myoblast fusion, at
16 h
after puparium formation (Fernandes et
al., 1991
).
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Phosphorylation of flightin begins during late stages of pupal development,
culminating in nine phosphovariants in mature adults
(Vigoreaux and Perry, 1994).
We conducted 2DE analysis to determine if the premature expression of
flightin in P{fln+}0.2 results in
premature phosphorylation, as is seen in some IFM mutants
(Vigoreaux, 1994
).
Fig. 4 shows the pattern of
phosphovariant accumulation in P{fln+}0.2 is
undistinguishable from that in wild-type flies from stage P15 through
adult.
|
Next we determined if levels of flightin expression are different in transgenic and wild-type strains. The relative abundance of flightin was estimated by western blot analysis after normalization to total thoracic protein (see Materials and methods). The rescued line showed a small reduction in flightin levels (Fig. 5).
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Normal fiber mechanics in wild-type flightin transgenic lines
The increased compliance of IFM fibers devoid of flightin results in loss
of power output, most likely due to internal absorption of a large amount of
the actomyosin generated work (Table
2) (Henkin et al.,
2004). There is no statistically significant difference in the
dynamic stiffness of fibers from P{fln+}0.2
rescued flies and those of wild-type flies. The complex modulus (an index of
dynamic stiffness) is composed of two components, the elastic modulus
(Ee), and the viscous modulus (Ev).
Ee is a measure of fiber compliance and in
P{fln+}0.2-rescued flies Ee
is statistically the same as wild type at the frequency at which maximum power
generation occurs (fmax)
(Table 2).
Fig. 6A is a plot of
Ee vs frequency at maximal Ca2+
activation (pCa 5). Note that P{fln+}0.2 produces
a normal triphasic response, indicative of restoration of wild-type function.
Ev is a measure of the work produced (negative values) and
absorbed (positive values) by the fiber.
Fig. 6B demonstrates that
Ev values for P{fln+}0.2 are
nearly identical to wild type and the value at the frequency at which maximum
power occurs is not statistically different from wild type
(Table 2). Power production by
P{fln+}0.2 fibers is lower than power production
by wild-type fibers but the differences are not statistically significant
(Fig. 6C and
Table 2).
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Increased flightin gene copy number does not have an effect on muscle structure or contractile properties
We studied one line of wild-type flies that had been transformed with
P{fln+} (w1118,
P{w+, Act88F-fln+};fln+, abbreviated
P{fln+}2.2) to determine if increasing gene copy
number has an effect on flightin protein levels and IFM properties. Flies that
carry four copies of the flightin gene have normal flight ability
(Table 1). The myofibrillar
structure is also normal (Fig.
2) with regular sarcomere length and a normal number of thick
filaments per myofibril (Table
1). Despite the doubling in flightin gene copy number,
P{fln+}2.2 flies do not exhibit an increase in
flightin protein accumulation relative to wild-type flies, but a significant
increase relative to P{fln+}0.2 rescued flies
(Fig. 5). In all respects,
P{fln+}2.2 are more similar to wild-type flies
than are P{fln+}0.2 flies.
Unlike flightin protein levels, expression of MHC protein increases with
doubling of gene copy number. Thus, flies carrying four copies of the
Mhc+ gene express 24 times more myosin than normal
diploid flies and also have an over-abundance of thick filaments
(Cripps et al., 1994). We
determined the expression levels of flightin in the Drosophila line
w; P{w+Mhc+}wm2 and found that it was not
significantly different from P{fln+}0.2 but
significantly less than wild type. Sinusoidal analysis of skinned
P{fln+}2.2 fibers revealed that their mechanical
properties are on par with wild-type fibers
(Table 2 and
Fig. 6).
Muscle power output in vivo decreases in flightin transgenic lines
While flying in the virtual reality flight simulator, flight performance of
the Drosophila transgenic rescued line
P{fln+}0.2 and the multi-gene copy line
P{fln+}2.2 is significantly reduced during
maximum locomotor capacity compared with wild-type flies. Although all flies
generate enough flight force to sustain hovering flight,
Table 3 shows that the reduced
capability of the transgenic lines to produce flight force in excess of
hovering flight force appears to be due to a reduction in both stroke
amplitude (P{fln+}2.2) and frequency
(P{fln+}0.2 and
P{fln+}2.2), whereas muscle and aerodynamic
efficiency appear to be widely similar in the three lines.
|
Aerodynamic flight force reduction amounts to 15% in
P{fln+}0.2 and 24% in
P{fln+}2.2 compared with wild type that is
correlated with a significant reduction in both stroke amplitude of
approximately 5 and 13 degrees, and stroke frequency of 23 and
9 Hz,
respectively. As a consequence, the cost of generating lift (induced power)
decreases significantly by
31 and
36% in the two transgenic lines
compared with the control animals. Similar results were obtained for profile
power, the cost to overcome the drag on the moving wings
(Table 3). Muscle mechanical
power output in the behaving flies, given as the sum of induced and profile
power, decreases in the transgenic lines by approximately 29%
(P{fln+}0.2) and 34%
(P{fln+}2.2) compared with wild type. A similar
trend was observed at the single fiber level. Power output by
P{fln+}0.2 and
P{fln+}2.2 fibers was decreased by 15% and 26%,
respectively, compared with wild type
(Table 2). However, these
differences were not statistically significant.
The reduction in muscle performance is consistent with a reduction in metabolic power, yielding constant values of muscle efficiency ranging from 8.89% in P{fln+}2.2 to 9.76% in wild type. The modification in flightin expression did not alter aerodynamic efficiency of force production between the lines that ranges from 25.7% to 26.8% indicating that the cost of flight force production due to wing flapping did not change among the three lines.
We did not find any significant differences in maximum flight force
production, muscle mechanical power output, metabolic power and the two
efficiency estimates between P{fln+}0.2 and
P{fln+}2.2. Interestingly, the two transgenic
lines generated maximum flight force using different combinations of stroke
amplitude and stroke frequency. It was shown previously that force production
in Drosophila linearly depends on wing velocity, given by the product
of amplitude and frequency (Lehmann and
Dickinson, 1998). Although mean wing velocity at the center of
wing area is similar in both transgenic Drosophila lines of
approximately 1.97±0.1 (P{fln+}0.2) and
2.01±0.2 m s-1 (P{fln+}2.2),
stroke amplitude was significantly higher in
(P{fln+}0.2) line compared with
P{fln+}2.2 whereas stroke frequency was
approximately 6% higher in P{fln+}2.2 compared
with (P{fln+}0.2). The latter is consistent with
results at the fiber level where fmax is 13% higher in
P{fln+}2.2 than in
P{fln+}0.2. The kinematic differences are
accompanied by small but notably significant differences in body mass (body
mass P{fln+}0.2=0.88 mg,
P{fln+}2.2=1.01 mg) and a small but significant
reduction in wing area between P{fln+}0.2 (1.87
mm2) and wild-type flies (2.00 mm2;
Table 3).
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Discussion |
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One remarkable feature of rescued P{fln+}0.2
IFM is the decreased number of thick filaments per sarcomere. This decrease is
not a result of flightin under-expression given that nearly normal
levels of the protein are found in the mature IFM. Furthermore, a mutation
that results in flightin under-expression shows a distinctly
different phenotype (see below). A more plausible explanation is that the
slightly premature expression of flightin during pupal development of
P{fln+}0.2 interferes with some aspect of thick
filament assembly. The Act88F promoter, which drives expression of
the flightin transgene, has been shown to be activated very early in
IFM development (16 h after puparium formation;
Fernandes et al., 1991
). While
this study did not examine the timing of transcriptional activation of the
Act88Ffln transgene, it is evident from western blot
analysis (Fig. 3) that the
transgene-encoded flightin protein begins to accumulate earlier than the
endogenous gene-encoded flightin in wild-type flies (pupal stage P7
vs P8, respectively). The temporal program of flightin
phosphorylation, however, seems to be unaffected by the untimely expression
(Fig. 4).
In a previous study we had shown that Drosophila heterozygous for
a deficiency that encompasses the flightin gene,
Df(3L)fln1, showed an 20% reduction in flightin
(Vigoreaux et al., 1998
). This
resulted in myofibrillar defects, evident as peripheral disassociation of the
thick and thin filaments, as well as altered fiber kinetics and attenuated
flight (Vigoreaux et al.,
1998
). Thus, while both P{fln+}0.2
and Df(3L)fln1 retain an intact myofibril core that is
80% the diameter of the intact myofibril, Df(3L)fln1
exhibited loosely organized peripheral myofilaments while
P{fln+}0.2 exhibited an
17% reduction in the
number of myofilaments. A second difference is that fiber power output is
reduced by
15% in P{fln+}0.2 but unchanged
in Df(3L)fln1 that instead exhibited an increase in
fmax. Altogether, these studies suggest that the premature
accumulation of flightin in P{fln+}0.2 is the
most likely explanation for the reduced number of thick filaments and that
unphosphorylated flightin participates in the process by which the number of
thick filaments is determined during sarcomerogenesis. One possibility is that
`premature' flightin binds monomeric myosin and prevents its incorporation
into a growing polymer, perhaps by interfering with the electrostatic
interactions between myosin rod coiled coils that are required for assembly
(Atkinson and Stewart, 1991
;
McLachlan and Karn, 1982
).
Future studies will investigate this possibility.
There is one other example where timing of expression adversely affects IFM
development. Transgenic flies that express an Act88F
promoter-mini-paramyosin chimeric gene in their IFM show subtle developmental
defects that are compounded in the adult working muscle
(Arredondo et al., 2001).
Unlike flightin, mini-paramyosin that is under Act88F
promoter regulation is over-expressed
(Arredondo et al., 2001
).
The reduced myofibrillar diameter in P{fln+}0.2 does not appear to have deleterious effect on fiber mechanics and flight parameters. Dynamic stiffness and power output from skinned P{fln+}0.2 fibers were more similar to wild type than the corresponding values from P{fln+}2.2, despite the fact that the latter had the normal number of thick filaments per sarcomere. Likewise, normalized force and mechanical power measured in the flight arena for P{fln+}0.2 are more similar to wild-type values than P{fln+}2.2 values are, as are muscle and aerodynamic efficiency. The differences between any of the above parameters for P{fln+}0.2 and P{fln+}2.2 are not statistically significant. However, the observation that all values follow a similar trend suggest that the presence of extra copies of the flightin gene, while restoring the quota of thick filaments, has a moderately unfavorable effect on flight muscle function.
Measurements in the flight arena also revealed large differences between
wild type and the two transgenic strains, differences that were not evident on
the mechanical analysis of skinned single IFM fibers. One interpretation, as
already surmised, is that P{fln+}0.2 is not fully
rescued while P{fln+}2.2 exhibits detrimental
effects of tetraploidy. It is not uncommon for transgenic strains not to
perform to the same level as wild-type strains. For example, the wild-type
Mhc gene can rescue the flightlessness imposed by the amorphic
Mhc10 allele but transgenic flies are not fully flighted
(Cripps et al., 1994).
Similarly, the wild-type Tropomyosin (Tm2) gene was able to
rescue the flightless behavior and IFM contractile defects engendered by the
Tm2 deletion allele TmIC10, but performance fell
short of that of wild-type flies (Kreuz et
al., 1996
). Thus under-performance of transgenic animals appears
to be a general feature in Drosophila perhaps as a result of the
random genomic integration of the P-element-shuttled transgene.
A second interpretation is that genetic differences among the strains, more
so than the ability of the transgene to rescue the mutant phenotype, accounts
for the variability. Allele differences among genes that directly or
indirectly affect flight behavior cannot be completely ruled out even among
strains that were derived from a common parental strain because their
generation required different outcrosses. For example, the
fln0-carrying chromosome is marked by an ebony
allele, a mutation that exhibits a variety of locomotor rhythm anomalies
although none that is known to affect flight. Yet a third interpretation is
that the differences in the flight arena reflect the contributions of muscles
other than the DLM, in particular those of the opposing set of IFM, the
dorsoventral muscles (DVM). It is assumed that the DVM have the same
contractile properties (and flightin expression) as the DLM but this
has not been experimentally tested given the greater difficulty of isolating
DVM fibers. One important difference is that the DLM develops from a scaffold
of larval muscles while the DVM develops de novo by fusion of
imaginal myoblasts (Fernandes et al.,
1991; for review see:
Vigoreaux and Swank, 2004
).
The different developmental pathways of DLM and DVM may impose distinct
regulatory constraints on the expression of the Act88Ffln
transgene resulting in greater functional differences among these two fiber
types.
IFM has been shown to be very sensitive to expression levels for a variety
of its constituent proteins (for review see
Vigoreaux and Swank, 2004). As
mentioned earlier, over-expression of mini-paramyosin resulted in flight
defects (Arredondo et al.,
2001
). Myofibril assembly occurred normally, and myofibrils in
young adults were relatively normal, but as flies aged, degeneration occurred
so that by 10 days into adulthood, there was considerable myofibrillar
degeneration that translated into severe flight impairment. Over-expression of
a heat shock-sanpodo (spdo) transgene, the Drosophila
Tropomodulin homolog, during mid-to-late pupal stages caused shorter than
normal thin filaments in IFM and flight impairment
(Mardahl-Dumesnil and Fowler,
2001
).
Mhc tetraploidy (P{w+Mhc+}wm2)
resulted in a twofold increase in myosin expression, excess and loosely
associated thick filaments residing in the myofibrillar peripheries, and a
severe flight defect (Cripps et al.,
1994). Some of the peripheral thick filaments also appeared to
have a smaller diameter than those in the center of the myofibril. Given the
excess thick filaments, it was surprising to find that flightin levels were
lower in P{w+Mhc+}wm2 than in wild type. One
possible explanation for this observation is that the excess myosin
outcompetes flightin for myosin binding during polymerization and unassembled
flightin is rapidly degraded. This scenario is consistent with our proposal
above that `premature' flightin binds and `hijacks' monomeric myosin,
resulting in less thick filaments polymerized. A second possibility is that
the absence of thin filaments and of a well formed lattice in the myofibril
periphery creates an environment where flightin is unstable. The absence of
thin filaments is known to affect accumulation of flightin phosphovariants
(Vigoreaux, 1994
). It is
interesting to note that the amorphous myofibril periphery in
P{w+Mhc+}wm2 is not unlike that seen in
Df(3L)fln1 heterozygotes.
In contrast to Mhc, but similar to P{fln+}2.2,
tetraploidy of Act88F does not result in loss of flight ability
(Hiromi et al., 1986). Because
the study relied only on a simple flight test, it is not possible to establish
if excess actin genes affected IFM function in ways that are not evident in
the flight test. A recent study showed that copy number polymorphism is rather
common in `normal' humans (Sebat et al.,
2004
). While some of the polymorphisms may be associated with
susceptibility to health problems, others may effect no phenotype. Hence
expression of particular proteins in humans, like in flies, is influenced by
gene copy number while expression of other proteins is not.
In summary, our results show that relative levels of flightin accumulation in the IFM are not strictly dictated by gene copy number, as has been demonstrated for other myofibrillar proteins. Instead, regulation of flightin levels appears to be tightly dependant on the process of thick filament and myofibril assembly, perhaps dictated by the availability of myosin binding sites and/or the integrity of the myofibrillar lattice. The results bring a new dimension to our understanding of myofibril assembly as they underscore the need to understand the role of protein interactions in addition to gene regulatory mechanisms. Proper regulation of flightin levels is essential for normal myofibrillogenesis and flight muscle function. Transgenic studies such as the one described here will continue to be pursued to further define the functional roles of flightin in muscle development and contraction.
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References |
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Arredondo, J. J., Mardahl-Dumesnil, M., Cripps, R. M., Cervera, M. and Bernstein, S. I. (2001). Overexpression of miniparamyosin causes muscle dysfunction and age-dependant myofibril degeneration in the indirect flight muscles of Drosophila melanogaster.J. Muscle Res. Cell Motil. 22,287 -299.[CrossRef][Medline]
Atkinson, S. J. and Stewart, M. (1991). Molecular basis of myosin assembly: coiled-coil interactions and the role of charged periodicities. J. Cell Sci. Suppl. 14, 7-10.[Medline]
Ayer, G. and Vigoreaux, J. O. (2003). Flightin is a myosin rod binding protein. Cell Biochem. Biophys. 38,41 -54.[Medline]
Bainbridge, S. P. and Bownes, M. (1981). Staging the metamorphosis of Drosophila melanogaster. J. Embryol. Exp. Morph. 66,57 -80.[Medline]
Bernstein, S. I., O'Donnell, P. T. and Cripps, R. M. (1993). Molecular genetic analysis of muscle development, structure and function in Drosophila. Int. Rev. Cytol. 143,63 -152.[Medline]
Cripps, R. M. (2004). The contributions of genetics to the study of insect flight muscle function. In Nature's Versatile Engine: Insect Flight Muscle Inside and Out, in press (ed. J. Vigoreaux). Georgetown, TX: Landes Bioscience.
Cripps, R. M., Becker, K. D., Mardahl, M., Kronert, W. A., Hodges, D. and Bernstein, S. I. (1994). Transformation of Drosophila melanogaster with the wild-type myosin heavy-chain gene: rescue of mutant phenotypes and analysis of defects caused by overexpression. J. Cell Biol. 126,689 -699.[Abstract]
Cripps, R. M., Suggs, J. A. and Bernstein, S. I.
(1999). Assembly of thick filaments and myofibrils occurs in the
absence of the myosin head. EMBO J.
18,1793
-1804.
Dickinson, M. H., Hyatt, C. J., Lehmann, F.-O., Moore, J. R., Reedy, M. C., Simcox, A., Tohtong, R., Vigoreaux, J. O., Yamashita, H. and Maughan, D. W. (1997). Phosphorylation-dependent power output of transgenic flies: an integrated study. Biophys. J. 73,3122 -3134.[Abstract]
Dickinson, M. H., Lehmann, F. O. and Sane, S. P.
(1999). Wing rotation and the aerodynamic basis of insect flight.
Science 284,1954
-1960.
Ellington, C. P. (1984). The aerodynamics of insect flight. IV. Aerodynamic mechanisms. Phil. Trans. R. Soc. Lond. B 305,79 -113.
Fernandes, J., Bate, M. and Vijayraghavan, K. (1991). Development of the indirect flight muscles of Drosophila. Development 113, 67-77.[Abstract]
Geyer, P. K. and Fyrberg, E. A. (1986). 5'-flanking sequence required for regulated expression of a muscle-specific Drosophila melanogaster actin gene. Mol. Cell Biol. 6,3388 -3396.[Medline]
Henkin, J. A., Maughan, D. W. and Vigoreaux, J. O.
(2004). Mutations that affect flightin expression in
Drosophila alter the viscoelastic properties of flight muscle fibers.
Am. J. Physiol. Cell Physiol.
286,C65
-C72.
Hiromi, Y., Okamoto, H., Gehring, W. J. and Hotta, Y. (1986). Germline transformation with Drosophila mutant actin genes induces constitutive expression of heat shock genes. Cell 44,293 -301.[CrossRef][Medline]
Klemenz, R., Weber, U. and Gehring, W. J. (1987). The white gene as a marker in a new P element vector for gene transfer in Drosophila. Nucleic Acids Res. 15,3947 -3959.[Abstract]
Kreuz, A. J., Simcox, A. and Maughan, D. (1996). Alterations in flight muscle ultrastructure and function in Drosophila tropomyosin mutants. J. Cell Biol. 135,673 -687.[Abstract]
Kronert, W. A., O'Donnell, P. T., Fieck, A., Lawn, A., Vigoreaux, J. O., Sparrow, J. C. and Bernstein, S. I. (1995). Defects in the Drosophila myosin rod permit sarcomere assembly but cause flight muscle degeneration. J. Mol. Biol. 249,111 -125.[CrossRef][Medline]
Laemmli, U. K. (1970). Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 270,680 -685.
Lehmann, F. O. and Dickinson, M. H. (1998). The
control of wing kinematics and flight forces in fruit flies
(Drosophila spp.). J. Exp. Biol.
201,385
-401.
Lehmann, F.-O. and Dickinson, M. H. (1997). The
changes in power requirements and muscle efficiency during elevated flight
force production in the fruit fly, Drosophila. J. Exp.
Biol. 200,1133
-1143.
Liu, H., Mardahl-Dumesnil, M., Sweeney, S. T., O'Kane, C. J. and
Bernstein, S. I. (2003). Drosophila paramyosin is
important for myoblast fusion and essential for myofibril formation.
J. Cell Biol. 160,899
-908.
Mardahl-Dumesnil, M. and Fowler, V. M. (2001).
Thin filaments elongate from their pointed ends during myofibril assembly in
Drosophila indirect flight muscle. J. Cell
Biol. 155,1043
-1053.
McLachlan, A. D. and Karn, J. (1982). Periodic charge distributions in the myosin rod amino acid sequence match cross-bridge spacings in muscle. Nature 299,226 -231.[CrossRef][Medline]
Miller, B. M. and Bernstein, S. I. (2004). Myosin. In Nature's Versatile Engine: Insect Flight Muscle Inside and Out, in press (ed. J. Vigoreaux). Georgetown, TX: Landes Bioscience.
Miller, R. C., Schaaf, R., Maughan, D. W. and Tansey, T. R. (1993). A non-flight muscle isoform of Drosophila tropomyosin rescues an indirect flight muscle tropomyosin mutant. J. Muscle Res. Cell Motil. 14, 85-98.[Medline]
Nongthomba, U. and Ramachandra, N. B. (1999). A
direct screen identifies new flight muscle mutants on the Drosophila
second chromosome. Genetics
153,261
-274.
Reedy, M. C. and Beall, C. (1993). Ultrastructure of developing flight muscle in Drosophila. I. Assembly of myofibrils. Dev. Biol. 160,443 -465.[CrossRef][Medline]
Reedy, M. C., Bullard, B. and Vigoreaux, J. O.
(2000). Flightin is essential for thick filament assembly and
sarcomere stability in Drosophila flight muscles. J. Cell
Biol. 151,1483
-1499.
Rubin, G. M. and Spradling, A. C. (1983). Vectors for P element-mediated gene transfer in Drosophila. Nucleic Acids Res. 11,6341 -6351.[Abstract]
Sanchez, F., Tobin, S. L., Rdest, U., Zulauf, E. and McCarthy, B. J. (1983). Two Drosophila actin genes in detail. Gene structure, protein structure and transcription during development. J. Mol. Biol. 163,533 -551.[Medline]
Sebat, J. et al. (2004). Large-scale copy
number polymorphism in the human genome. Science
305,525
-528.
Spradling, A. C. and Rubin, G. M. (1982). Transposition of cloned P elements into Drosophila germ-line chromosomes. Science 218,341 -347.[Medline]
Swank, D. M., Wells, L., Kronert, W. A., Morrill, G. E. and Bernstein, S. I. (2000). Determining structure/function relationships for sarcomeric myosin heavy chain by genetic and transgenic manipulation of Drosophila. Micro. Res. Tech. 50,430 -442.[CrossRef]
Vigoreaux, J. O. (1994). Alterations in flightin phosphorylation in Drosophila flight muscles are associated with myofibrillar defects engendered by actin and myosin heavy chain mutant alleles. Biochem. Genet. 32,301 -314.[CrossRef][Medline]
Vigoreaux, J. O. (2001). Genetics of the Drosophila flight muscle myofibril: a window into the biology of complex systems. BioEssays 23,1047 -1063.[CrossRef][Medline]
Vigoreaux, J. O. and Perry, L. M. (1994). Multiple isoelectric variants of flightin in Drosophila stretch-activated muscles are generated by temporally regulated phosphorylations. J. Muscle Res. Cell Motil. 15,607 -616.[Medline]
Vigoreaux, J. O. and Swank, D. M. (2004). The development of the flight and leg muscle. In Comprehensive Molecular Insect Science, vol. 2 (eds. L. I. Gilbert, K. Iatrou and S. Gill), pp. 45-84. Oxford, UK: Elsevier.
Vigoreaux, J. O., Saide, J. D. and Pardue, M. L. (1991). Structurally different Drosophila striated muscles utilize distinct variants of Z-band-associated proteins. J. Muscle Res. Cell Motil. 12,340 -354.[Medline]
Vigoreaux, J. O., Saide, J. D., Valgeirsdottir, K. and Pardue, M. L. (1993). Flightin, a novel myofibrillar protein of Drosophila stretch-activated muscles. J. Cell Biol. 121,587 -598.[Abstract]
Vigoreaux, J. O., Hernandez, C., Moore, J., Ayer, G. and
Maughan, D. (1998). A genetic deficiency that spans the
flightin gene of Drosophila melanogaster affects the ultrastructure
and function of the flight muscles. J. Exp. Biol.
201,2033
-2044.