Acidic and basic troponin T isoforms in mature fast-twitch
skeletal muscle and effect on contractility
Ozgur
Ogut1,
Henk
Granzier2, and
Jian-Ping
Jin1
1 Department of Physiology and
Biophysics, Case Western Reserve University School of Medicine,
Cleveland, Ohio 44106-4970; and
2 Department of Veterinary and
Comparative Anatomy, Pharmacology, and Physiology, Washington State
University, Pullman, Washington 99164-6520
 |
ABSTRACT |
Developmentally
regulated alternative RNA splicing generates distinct classes of acidic
and basic troponin T (TnT) isoforms. In fast-twitch skeletal muscles,
an acidic-to-basic TnT isoform switch ensures basic isoform expression
in the adult. As an exception, an acidic segment in the
NH2-terminal variable region of
adult chicken breast muscle TnT isoforms is responsible for the unique exclusive expression of acidic TnTs in this muscle (O. Ogut and J.-P.
Jin. J. Biol. Chem. 273:
27858-27866, 1998). To understand the relationship between acidic
vs. basic TnT isoform expression and muscle contraction, the
contractile properties of fibers from adult chicken breast muscle were
compared with those of the levator coccygeus muscle, which expresses
solely basic TnT isoforms. With use of Triton X-100-skinned muscle
fibers, the force and stiffness responses to
Ca2+ were measured. Relative to
the levator coccygeus muscle, the breast muscle fibers showed
significantly increased sensitivity to
Ca2+ of force and stiffness with a
shift of ~0.15 in the pCa at which force or stiffness was 50% of
maximal. The expression of tropomyosin, troponin I, and troponin C
isoforms was also determined to delineate their contribution to
thin-filament regulation. The data indicate that TnT isoforms differing
in their NH2-terminal charge are
able to alter the sensitivity of the myofibrillar contractile apparatus to Ca2+. These results provide
evidence linking the regulated expression of distinct acidic and basic
TnT isoform classes to the contractility of striated muscle.
alternative ribonucleic acid splicing; developmental regulation; calcium; activation of force and stiffness; tropomyosin
 |
INTRODUCTION |
TROPONIN T (TnT) is the tropomyosin
(Tm)-binding subunit of the troponin complex and a central element in
the thin-filament-linked Ca2+
regulatory system of vertebrate striated muscle (20). A large diversity
of TnT isoforms is expressed in striated muscles as a result of
alternative RNA splicing (for recent review see Ref. 31). The
5'-variable region of the TnT transcript is responsible for
multiple isoforms that differ in their
NH2-terminal primary structure (3,
7, 14, 19, 35, 36, 40). In addition, alternative splicing of the
mutually exclusive exons 16 and 17 generates additional isoforms from
the fast-twitch skeletal muscle TnT gene (3, 36, 40, 43). Although the
alternatively spliced NH2-terminal
variable region (13, 29) is quantitatively acidic in all TnTs, the
expression of alternatively spliced exons results in TnT isoforms with
a wide range of overall
NH2-terminal charge. A common
theme in the pattern of TnT isoform expression during cardiac and
skeletal muscle development is the regulated high-to-low molecular
weight (Mr) and
acidic-to-basic isoform switch (16, 40). The functional significance of
the switch between TnT isoform classes of distinct physical properties
remains largely unknown because of the isoform diversity, which
complicates the characterization of individual TnTs. Nonetheless,
differences in the Ca2+
sensitivity of the actomyosin-ATPase were demonstrated in reconstituted systems containing two bovine cardiac TnT isoforms with differences in
NH2-terminal size and charge (39).
Studies have correlated TnT isoform expression with muscle
contractility in normal and pathological states (1, 2, 34), although
these investigations have not delineated the physical properties of the
TnT isoforms that change in expression level. While the
NH2-terminal variable region
appears to be nonessential for TnT's core function in actomyosin activation (28), we have demonstrated that the
NH2-terminal structure is able to
modulate TnT's conformation and interaction with other thin-filament
proteins (25, 41). Furthermore, acidic and basic fast-twitch skeletal
muscle TnT isoforms have differences in their ability to bind Tm and
troponin I (TnI) in response to decreased pH, indicating that the
NH2 terminus of TnT may contribute to the tolerance of muscle to acidosis (26).
The physiological significance of acidic and basic TnT isoforms needs
to be further characterized in an integrated muscle system. In the
present study we have investigated the relationship between the
expression of acidic and basic fast TnT isoforms and the contractility
of muscle. Using skinned fibers from adult chicken breast muscle
(exclusive acidic TnT expression) and levator coccygeus (exclusive
basic TnT expression), we show that acidic TnT expression contributed
to the function of the contractile apparatus by sensitizing force and
stiffness responses to Ca2+.
Therefore, the increased expression of basic TnT isoforms in adult
muscle may contribute to the lower sensitivity to
Ca2+ than in neonatal muscle (37),
implying an important modulatory role for TnT isoforms in the fine
tuning of muscle contraction.
 |
MATERIALS AND METHODS |
Preparation of muscle homogenates.
Adult White Leghorn chicken (Gallus
domesticus) muscles were identified according to
Nickel et al. (24) and excised. A sample (~100 mg) of the fresh
muscle was immediately homogenized in 1 ml of SDS-PAGE sample buffer
containing 1% SDS and heated to 80°C for 5 min. The total protein
extracts were clarified by centrifugation at 14,000 g for 5 min in a microcentrifuge
before SDS-PAGE.
Western blot analysis of TnT, Tm, and TnI isoform expression.
The 1% SDS extracts of muscles were resolved by two SDS-PAGE systems
to maximize the resolution of protein isoforms:
1) 12% Laemmli SDS-PAGE with an
acrylamide-to-bisacrylamide ratio of 29:1 or
2) 14% Laemmli SDS-PAGE with an
acrylamide-to-bisacrylamide ratio of 180:1. In 29:1 12% SDS-PAGE,
low-Mr TnT
isoforms are well separated, but
high-Mr TnT
isoforms comigrate with actin, causing a broadening of these TnT bands
in Western blots. In 180:1 14% SDS-PAGE,
high-Mr TnT
isoforms are well resolved and separated from the actin band, but
stacking of
low-Mr TnT
isoforms may occur. Resolved proteins were transferred to 0.45-µm
nitrocellulose membranes, as previously described (25). Replica
nitrocellulose membranes were incubated at 4°C overnight with
rabbit polyclonal antisera raised against chicken breast muscle TnT
(RATnT) (41) or a chicken fast TnT
NH2-terminal peptide (Tx)
conjugated to keyhole limpet hemocyanin (RATx) (18), an anti-Tx
monoclonal antibody (MAb) 6B8 (41), a chicken fast-twitch skeletal
muscle TnT-specific MAb 3E4 (41), a cardiac muscle TnT-specific MAb CT3
(17), an
- and
-isoform-specific anti-Tm MAb CH1 (22) (provided by Dr. J. J.-C. Lin, University of Iowa), or a TnI-specific MAb TnI-1
(J.-P. Jin and F. Yang, unpublished results). The subsequent washing,
incubation with alkaline phosphatase-labeled anti-mouse or rabbit IgG
secondary antibody (Sigma Chemical), and
5-bromo-4-chloro-3-indolylphosphate-nitro blue tetrazolium color
development were performed as previously described (25).
Purification and identification of troponin C.
To determine troponin C (TnC) isoform expression, a recombinant
prokaryotic expression plasmid encoding chicken fast-twitch skeletal
muscle TnC (kindly provided by Dr. L. B. Smillie, University of
Alberta) was used to express TnC in Escherichia
coli BL21(DE3)pLysS. The culture medium, growth, and
induction conditions have been previously described (25). The culture
was induced by 0.2 mM isopropyl-1-thio-
-D-galactopyranoside
at an optical density at 600 nm of 0.9 and grown for another 3 h. After
the induced bacterial culture was harvested, the cells were resuspended
in 6 M urea, 30 mM Tris · HCl, pH 8.0, and 2 mM
MgCl2 and lysed by three passes through a French press at 500-700 psi. The clarified lysate was loaded onto a DE52 ion-exchange column equilibrated in the same buffer
and eluted by a 0-300 mM KCl gradient. The fractions containing the TnC peak were identified by SDS-PAGE, collected, and dialyzed for
two changes against 4 liters of double-distilled water. After lyophilization the protein powder was resuspended in a minimal volume
of 0.5 M KCl, 20 mM Tris · HCl, pH 8.0, and 2 mM
MgCl2 and resolved by a gel
filtration column (Sephadex G75, Pharmacia-Amersham). The TnC peak from
the gel filtration column was identified and dialyzed as described
above before lyophilization for long-term storage at
20°C.
With the purified chicken fast-twitch skeletal muscle TnC as an
immunogen, a mouse antiserum (MATnC) was raised and used to identify
TnC isoforms in Western blots of fiber homogenates.
To provide a native fast-twitch skeletal muscle TnC control, TnC was
also purified from chicken breast muscle, as described by Potter (32).
To monitor the difference in migration between the skeletal and cardiac
TnC isoforms, a prokaryotically expressed mouse cardiac TnC protein was
used. E. coli BL21(DE3)pLysS was transformed with a pET3d (Novagen) expression plasmid encoding mouse
cardiac TnC (A. Chen and J.-P. Jin, unpublished results), cultured, and
induced with
isopropyl-1-thio-
-D-galactopyranoside, as described above, to express cardiac TnC. Total protein extracts from
the bacterial cultures were prepared for SDS-PAGE, as described previously (25). To maximize the separation of TnC isoforms by
SDS-PAGE, 15% Laemmli gel with an acrylamide-to-bisacrylamide ratio of
29:1 was used.
Preparation of skinned chicken muscle fibers.
Fiber bundles were dissected from the pectoralis major and levator
coccygeus muscles of adult White Leghorn chickens after their
euthanization by CO2 inhalation.
The bundles were skinned for 60 min at room temperature in relaxing
solution [40 mM imidazole, 10 mM
bis-(aminoethyl)glycolether-N,N,N',N'-tetraacetic
acid, 6.4 mM magnesium acetate, 5.9 mM ATP sodium salt, 5 mM
NaN3, 80 mM potassium proprionate,
10 mM creatine phosphate, 1 mM dithiothreitol, 0.04 mM leupeptin, 0.5 mM phenylmethylsulfonate, pH 7.0] that contained 1% (wt/vol)
Triton X-100 (catalog no. 28314, Pierce Chemical). The bundles were
then washed twice with relaxing solution, twice with relaxing solution
in 50% glycerol and then stored at
20°C in relaxing
solution in 50% glycerol. The bundles were used within 2 wk.
Mechanical measurements.
The computer-controlled mechanics workstation used in this study has
been described in detail by Granzier and Irving (11). Briefly, a small
100-µl chamber was mounted on an x-y
stage of an inverted microscope that also contained a servomotor (model 6800, Cambridge Technology; step response ~0.3 ms, root mean
square position noise ~0.5 µm) and a force transducer
(model AME 801 E, Horton) with a strain gauge-conditioning amplifier
(model 2310, Measurement Group, Raleigh, NC) that had a bandwidth of
direct current of 10 kHz and a force resolution of ~100 µg. In
addition to force, high-frequency stiffness was measured using 0.1%
amplitude sinusoidal oscillations at 2.2 kHz. Force and stiffness
results were expressed per unit cross-sectional area. The
cross-sectional areas of the fibers were calculated from their measured
maximal and minimal diameters, with the assumption of an elliptical
cross section (11).
Single fibers were dissected from fiber bundles in relaxing
solution-50% glycerol. The fibers were then transferred to the microscope, and the ends were wrapped around fine pins (100 µm diameter) that had been glued to the motor and the force transducer. To
limit stretching of the wrapped portion of the fibers during a
contraction, a small volume (<1 µl) of 2% glutaraldehyde was added
to the wrapped portion of the fiber at the back side of the pin (5).
The fiber was quickly immersed into the chamber that was being rapidly
flushed with relaxing solution. The chamber was connected to a system
allowing continual perfusion of the muscle fibers with relaxing
solution or activating solution (pH of solutions = 7.0). The pCa of the
perfusing solution was varied by mixing relaxing solution with various
amounts of activating solution that contained the same components as
the relaxing solution, with the addition of
Ca2+ to 10 mM (8). In these
experiments the filament lattice spacing was not controlled. However,
no changes in fiber diameter were noticed as the pCa of the perfusing
solution was varied. The chamber contained a small J-type thermocouple
and was temperature controlled to 22°C in all experiments.
Sarcomere length measurement.
Sarcomere length was measured with laser diffraction with use of an
He-Ne laser beam focused to a diameter of ~250 µm. The diffraction
pattern was collected with a bright-field objective (ELWD plan 40/0.55,
Nikon); a telescope lens was focused on the back focal plane of the
objective, and the diffraction was projected, after compression with a
cylindrical lens, onto a photodiode array (model RL 256 C/17, Reticon).
The first-order diffraction peak position was obtained (12) using a
digital spot-position detector board (Dept. of Bioengineering,
University of Washington, Seattle, WA) installed in an IBM AT computer.
This signal was converted to sarcomere length by using a calibration
curve that was established with the diffraction peaks of a 25-µm
grating present in the chamber. Sarcomere length noise (peak to peak)
was ~10 nm. Sarcomere length was measured in the central region of
the muscle fibers and was 2.23 ± 0.10 and 2.28 ± 0.06 (SD) µm
during the steady force plateau of contractions of the pectoralis major
(n = 26) and levator coccygeus (n = 40) fibers, respectively.
After the mechanical measurements, cross sections from the fiber
bundles were homogenized in SDS-PAGE sample buffer, resolved by
SDS-PAGE, and immunoblotted using RATnT, 6B8, TnI-1, MATnC, and CH1
antibodies to verify the thin-filament protein isoforms in the fibers
of chicken pectoralis and levator coccygeus muscles.
 |
RESULTS |
Identification of thin-filament protein isoforms in adult chicken
muscles.
Using a panel of specific polyclonal and monoclonal antibodies, we
identified the TnT, TnI, TnC, and Tm isoforms expressed in
representative adult chicken striated muscles. The expression patterns
of fast-twitch skeletal muscle-specific (breast and gastrocnemius), slow-twitch skeletal muscle-specific (trapezius), and cardiac muscle-specific (left ventricle) thin-filament regulatory proteins were
determined (Fig. 1). In
fast-twitch skeletal muscle, immunoblots with RATnT show that a
heterogeneity of fast-twitch skeletal muscle TnT isoforms is expressed,
although these are present as two groups that differ by
Mr. MAb 6B8
staining for the acidic
NH2-terminal Tx segment indicates
that the high-Mr
isoform had an acidic NH2 terminus
compared with the
low-Mr
counterparts, which are the normal basic adult isoforms (26, 40). In
contrast, a single TnT isoform is identified by RATnT in the
slow-twitch trapezius muscle. Western blotting with the anti-cardiac
TnT MAb CT3, which cross-reacts with slow- but not fast-twitch skeletal
muscle TnT (14), indicates that the only TnT isoform expressed in
trapezius muscle is slow-twitch skeletal muscle TnT (44).
Interestingly, the RATnT antiserum generated against chicken breast
muscle TnT cross-reacts less with slow-twitch skeletal muscle TnT than
with cardiac muscle TnT (Fig. 1), given that comparable amounts of muscle protein extracts were loaded as normalized by the actin bands.
This indicates the presence of unique epitopes and the structural
divergence of slow-twitch skeletal muscle TnT vs. its fast-twitch
skeletal and cardiac muscle counterparts. Fast-twitch (lower
Mr) and
slow-twitch (higher
Mr) skeletal
muscle TnI isoforms were identified in the gastrocnemius and trapezius,
respectively. A single cardiac muscle TnI isoform was expressed in the
adult heart, and its migration was similar to slow-twitch skeletal
muscle TnI in the 180:1 14% gel. In contrast, SDS-PAGE of mammalian
cardiac and slow-twitch skeletal muscle TnI isoforms show significant differences in their apparent Mr (42). Tm
isoforms were mixed
- and
-Tm in the gastrocnemius and trapezius
muscles but exclusively
-Tm in the heart and the breast muscles.

View larger version (39K):
[in this window]
[in a new window]
|
Fig. 1.
Western blot identification of troponin T (TnT), tropomyosin (Tm), and
troponin I (TnI) isoforms. Specific antibodies were
used to determine TnT, Tm, and TnI isoforms expressed in adult chicken
striated muscles. Shown together with SDS-PAGE of comparatively loaded
samples, a variety of TnT isoforms are detected by a rabbit polyclonal
antiserum raised against chicken breast muscle TnT (RATnT). CT3
monoclonal antibody (MAb) staining shows that, in trapezius and heart,
only slow-twitch skeletal and cardiac muscle TnT isoforms are
expressed, respectively. Multiple fast-twitch skeletal muscle TnT
(fTnT) isoforms are present in fast-twitch skeletal muscles, as
detected by MAb 3E4 in breast and gastrocnemius muscles. Anti-Tx MAb
6B8 was used to specifically detect expression of an
NH2-terminal acidic stretch in
breast muscle TnT. The - and -isoforms of Tm were found, with
breast muscle and heart expressing exclusively -Tm. The fast-twitch
skeletal muscle isoform of TnI (fTnI) was found in breast and
gastrocnemius muscles, whereas slow-twitch skeletal muscle isoform TnI
(sTnI) was expressed in trapezius and cardiac isoform TnI (cTnI) was
expressed in heart. Gel concentration and acrylamide-to-bisacrylamide
ratio used for SDS-PAGE are indicated.
Mr, molecular
weight (relative).
|
|
With use of the MATnC antiserum raised against chicken fast-twitch
skeletal muscle TnC, TnC isoform expression was determined in cardiac
and fast- and slow-twitch skeletal muscles (Fig.
2). The chicken breast and the levator
coccygeus fibers expressed fast-twitch skeletal muscle TnC, indicated
by comigration with fast-twitch skeletal muscle TnC expressed from the
cloned cDNA or purified from chicken breast muscle. In contrast, the
slow-twitch trapezius muscle and cardiac muscle expressed TnC isoforms
of reduced mobility compared with fast-twitch skeletal muscle TnC. The
chicken slow-twitch/cardiac muscle TnC migrates similarly to the mouse
cardiac muscle TnC expressed from the cloned cDNA.

View larger version (48K):
[in this window]
[in a new window]
|
Fig. 2.
Troponin C (TnC) isoform expression in striated
muscles. A mouse antiserum raised against chicken
breast TnC (MATnC) was used to determine TnC isoform expression in
representative chicken striated muscles. With prokaryotically expressed
or natively purified chicken breast muscle TnC used as controls,
chicken breast and levator total homogenates are shown to express
fast-twitch skeletal muscle TnC. Trapezius and cardiac muscles
expressed
higher-Mr TnCs,
which migrated similarly to prokaryotically expressed mouse cardiac
TnC.
|
|
The expression patterns of acidic and basic fast-twitch skeletal muscle
TnT isoforms were determined by Western blotting with the RATnT and
RATx antisera (Fig. 3). The inclusion of
the Tx segment results in TnT isoforms with relatively acidic
isoelectric points and higher
Mr than their
basic counterparts (Fig. 1) (26). The majority of the muscles in the
pelvic limb of the chicken express basic TnT isoforms, consistent with
fast TnT isoforms identified in other adult skeletal muscles (40).
Therefore, the expression of acidic fast TnT isoforms in the adult
chicken breast muscles provides a novel model to study the
structure-function relationships of TnT.

View larger version (74K):
[in this window]
[in a new window]
|
Fig. 3.
Distribution of acidic and basic fast TnT isoforms in adult chicken
skeletal muscles. On total tissue homogenates from selected chicken
skeletal muscles, patterns of total TnT isoform expression and acidic
TnT isoform distribution were determined, respectively, by SDS-PAGE
(12%, 29:1) and Western blotting with use of RATnT and RATx antisera.
High-Mr, acidic
TnTs (widened bands were due to overlap with actin band) were
specifically expressed in pectoral limb muscles together with various
amounts of basic fast-twitch skeletal muscle TnT isoforms. A notable
exception was pectoralis superficialis (major), which expressed
exclusively
high-Mr, acidic
fast-twitch skeletal muscle TnT isoforms. Conversely, the majority of
muscles in the pelvic limb expressed
low-Mr, basic TnT
isoforms.
|
|
Developmental regulation of Tm expression.
In contrast to other fast-twitch skeletal muscles, the adult chicken
breast muscle expresses exclusively
-Tm. To determine whether the Tm
expression pattern in the breast muscle is developmentally regulated,
the Tm isoforms expressed in embryonic and adult breast major,
gastrocnemius, and heart muscles were identified (Fig. 4). The gastrocnemius and heart show
persistent mixed
- and
-Tm and
-Tm expression, respectively,
through development. Tropomyosin expression was mixed
and
isoforms in the embryonic breast muscle, but unlike other fast-twitch
skeletal muscles, the
-Tm isoform was downregulated in favor of the
exclusive expression of
-Tm in the adult. The developmental
regulation of Tm isoforms in chicken breast muscle shows a
correspondence to the increasing expression of acidic fast TnT isoforms
in breast muscle during development (26).

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 4.
Developmental regulation of Tm isoforms in developing fast-twitch
skeletal and cardiac muscles. Developmental expression
of Tm was determined in breast major, gastrocnemius, and heart by
Western blotting with use of 12% 29:1 SDS-PAGE on embryonic
day 14 (ED14) and adult muscle protein
extracts. Embryonic breast muscle expressed a mixture of - and
-Tm, similar to embryonic gastrocnemius, but -Tm expression was
downregulated with development in favor of -Tm. In contrast, there
was no change in Tm expression pattern in gastrocnemius through
development. Heart showed exclusive -Tm expression from embryo to
adult.
|
|
Ca2+
sensitivity of muscle fibers containing acidic or basic fast-twitch
skeletal muscle TnT isoforms.
To examine the relationship between TnT isoform expression and
Ca2+ sensitivity, mechanical
measurements were done with two representative, classical fast-twitch
white skeletal muscles: the pectoralis major and the levator coccygeus.
Both muscles produced high levels of isometric force in response to
perfusion with activating solution (Fig.
5). The performance of the fibers was
stable, and typically >10 contractions could be induced before a
noticeable force decrease took place. To determine how much of the ends
of the fibers had been fixed by glutaraldehyde, some of the fibers were
activated at the end of the experiment and allowed to highly shorten by moving the motor closer to the force transducer. All sarcomeres were
observed to shorten except those in a small region (~0.05-0.1 mm
long) at each end of the fiber that had been fixed by glutaraldehyde. The fixed ends greatly limited end compliance. Because of end compliance, sarcomeres in the central region of fibers with unfixed ends may significantly shorten during activation, even though the
length of the fiber is held constant; furthermore, the diffraction pattern of these sarcomeres often completely disappears. During our
experiments, however, the diffraction pattern of fibers with ends that
had been fixed remained very strong during activation and revealed that
sarcomeres in the central region of the fiber shortened only a very
short distance on activation: 43 ± 60 and 20 ± 43 (SD) nm for
pectoralis major (n = 26) and levator
coccygeus (n = 40), respectively.

View larger version (32K):
[in this window]
[in a new window]
|
Fig. 5.
Contractile response of skinned pectoralis major and levator coccygeus
(cocc) fibers. Representative tracings of response of sarcomere length,
force, and stiffness of skinned pectoralis and levator fibers to
Ca2+ at pCa 6.70 and 6.00 are
shown. Higher response of pectoralis than of levator fibers at pCa 6.7 indicates higher Ca2+ sensitivity
for force and stiffness.
|
|
The response of force and stiffness of the skinned pectoralis and
levator muscle fibers to Ca2+ were
measured. The Ca2+ sensitivities
of force (P < 0.02) and stiffness
(P < 0.01) were significantly higher
in the pectoralis than in the levator fibers, as judged by the
Ca2+ concentration required to
achieve 50% of maximal response (Figs. 6
and 7, Table 1). There was no significant
difference in the maximum force or stiffness per cross-sectional area
generated by the pectoralis or levator coccygeus fibers,
implying little difference in the maximum amount of cross bridges
formed or the force per cross bridge, as estimated by dividing maximal
force by maximal stiffness. Therefore, the
Ca2+ sensitivity of force and
stiffness in these muscles is reflective of regulation at the thin
filament. Examination of the slopes of the Hill plots in Figs. 6 and 7
(Table 1) showed slightly lower cooperativity of
Ca2+-activated force and stiffness
development in the pectoralis fibers, although the differences were not
statistically significant. It is interesting to note that stiffness
exhibits greater Ca2+ sensitivity
than force in pectoralis and levator fibers. A similar observation has
been reported by Martyn and Gordon (23), who investigated force and
stiffness in rabbit psoas fibers. Our results are consistent with their
proposal that attached cross bridges exist in non-force-producing and
force-producing states and that the relative population of these states
is Ca2+ dependent. At
low-to-intermediate Ca2+ levels
the non-force-producing attached population is relatively large,
explaining the high stiffness but low force levels.

View larger version (18K):
[in this window]
[in a new window]
|
Fig. 6.
Force-pCa relationship of skinned pectoralis major and levator
coccygeus muscle fibers. Pooled results and Hill fits of pooled data
are shown. Insets: Hill coefficients
and pCa at which force was 50% of maximal
(pCa50); results were obtained
from fitting each preparation separately and calculating mean ± SD
of 4 pectoralis major and 5 levator coccygeus fibers. Although the
lower Hill coefficient of pectoralis muscle was not statistically
significant, pCa50 of force and
stiffness are significantly higher for pectoralis than for levator
coccygeus fibers. Maximal force at pCa 6.0 was 160 ± 30 kN/m2 for pectoralis major and 130 ± 14 kN/m2 for levator
coccygeus fibers.
|
|

View larger version (20K):
[in this window]
[in a new window]
|
Fig. 7.
Stiffness response of skinned pectoralis major and levator coccygeus
muscle fibers. Pooled results and Hill fits from response of muscle
stiffness to free Ca2+ for
pectoralis major and levator coccygeus are shown. Pectoralis major
fibers showed higher sensitivity of stiffness to free
Ca2+, although maximal stiffness
at pCa 6.0 of the 2 muscles was not different: 13.2 ± 7.1 mN/m2 for pectoralis major and
13.4 ± 2.6 mN/m2 for levator
coccygeus fibers.
|
|
The thin-filament protein isoforms in the muscle fibers prepared for
contractility experiments were identified by immunoblotting with RATnT,
6B8, TnI-1, MATnC, and CH1 antibodies. The main difference between the
two muscles was the exclusive expression of
high-Mr acidic
TnTs in the pectoralis fibers and the exclusive expression of
low-Mr basic TnTs
in the levator fibers (Fig. 8).
Furthermore, the levator fibers showed heterogenous expression of
-
(major) and
(minor)-Tm isoforms, whereas the pectoralis fibers
showed predominantly
-Tm expression. TnI isoform expression was
comparable, with a trace amount of slow-twitch skeletal muscle TnI
expressed in the levator fibers. Both muscle fibers expressed
fast-twitch skeletal muscle TnC.

View larger version (18K):
[in this window]
[in a new window]
|
Fig. 8.
Thin-filament protein expression in skinned pectoralis and levator
coccygeus fibers. Muscle samples used for skinned fiber experiments
were homogenized in SDS-PAGE sample buffer and analyzed by Western
blotting. To compare the 2 muscle samples, nitrocellulose filters were
incubated with RATnT, 6B8, TnI-1, MATnC, or CH1 antibody to examine
expression of TnT, TnI, TnC, and Tm isoforms. Levator coccygeus fibers
showed expression of some -Tm and a trace amount of slow-twitch (S)
skeletal muscle TnI in addition to -Tm and fast-twitch (F) skeletal
muscle TnI. TnC isoform expression was identical in both muscles.
Cross-reactive bands, possibly myosin light chains, were also stained
by polyclonal anti-TnC serum, likely because of structural homology to
TnC (6). Pectoralis fibers expressed exclusively
high-Mr, acidic
fast-twitch skeletal muscle TnTs, whereas levator expressed
low-Mr, basic
fast-twitch skeletal muscle TnTs, accounting for the major difference
between the 2 fibers.
|
|
 |
DISCUSSION |
Unique expression of acidic TnT isoforms in adult chicken
fast-twitch skeletal muscle.
The developmental acidic-to-basic TnT isoform expression switch is
found in avian and mammalian cardiac and skeletal muscles. This
observed shift is most dramatic in fast-twitch skeletal muscles, where
the isoelectric points of the expressed TnT isoforms may increase from
5.04 in the neonate to 10.06 in the adult (
= 5.02) (40). The
importance of this large shift is not understood, since mouse cardiac
muscle TnT isoform expression patterns show relatively modest shifts in
their isoelectric point (
= 0.27) (19). Our survey of adult chicken
skeletal muscles demonstrated a heterogeneity of acidic and basic TnT
expression. As shown previously, high-Mr
fast-twitch skeletal muscle TnT isoforms in the mature chicken
(pectoralis muscles) correspond to isoforms that are acidic compared
with their low-Mr
counterparts (26). The expression of acidic fast-twitch skeletal muscle
TnTs in the adult chicken is unique, since all vertebrate animals
examined express exclusively basic fast-twitch skeletal muscle TnT
isoforms in adulthood (29, 40, 43). In chicken breast muscle the
inclusion of an acidic NH2-terminal segment results in
TnTs with significantly decreased isoelectric points (average 7.26)
compared with those in the gastrocnemius muscle (average 8.47) (26).
Although this segment is specific to the
Galliformes pectoral muscles (18), its
effect on TnT NH2-terminal size
and charge is representative of the difference between TnT isoform
classes during the acidic-to-basic switch in vertebrate skeletal muscle
(26, 40). Therefore, chicken breast muscle may serve as a model system
to determine the effects of acidic TnT isoform expression in the adult.
Effect of acidic fast-twitch skeletal muscle TnT isoforms on
thin-filament
Ca2+
sensitivity.
The two fast-twitch skeletal muscles tested showed differences in the
Ca2+ sensitivity of active force
and stiffness. Among the thin-filament regulatory proteins, fast-twitch
skeletal muscle TnI and TnC are expressed in both pectoralis and
levator coccygeus fibers (Fig. 8). Apart from the acidic vs. basic TnT
isoform expression, the two muscle types had differences in Tm isoform
expression, with the pectoralis fibers expressing only
-Tm and the
levator fibers expressing some
-Tm in addition to
-Tm (Fig. 8).
In previous studies, overexpression of
-Tm relative to
-Tm in
transgenic mouse hearts resulted in a slight sensitization of the force
response to Ca2+ in skinned
trabeculae (27). Because the levator coccygeus fibers show higher
expression of
-Tm but lower
Ca2+ sensitivity for force and
stiffness, Tm isoform expression cannot account for the difference in
Ca2+ sensitivity. In agreement
with this evidence, Reiser and co-workers (33) showed that although the
majority of pectoralis single fibers expressed exclusively
-Tm, a
minority of fibers expressed some
-Tm in addition to
-Tm.
Nonetheless, they found no differences in the
Ca2+ sensitivity of force between
these groups of fibers. Therefore, Tm isoform expression levels seem to
play a minor role in the responses of the fibers in these mechanical
measurements. We conclude that the differences in
Ca2+ sensitivity between the
levator and pectoralis fibers are likely due to TnT isoform expression,
with higher Ca2+ sensitivity in
muscles expressing acidic TnTs. This conclusion is consistent with
observations that the Ca2+
response of force is more sensitive in neonatal than in adult striated
muscles (10, 37), given that neonatal muscles express more acidic TnT
isoforms. Therefore, the acidic-to-basic TnT isoform switch in
developing avian and mammalian striated muscles (16, 19, 40) may
contribute to a change in the Ca2+
sensitivity of thin-filament activation and the overall cooperativity of muscle contraction (Fig. 6).
It is known that
-Tm is usually the exclusive Tm isoform found in
cardiac muscle (21), coexpressed with cardiac TnT, the most acidic of
all TnT isoforms. It was shown that
-Tm is downregulated during
development of the breast muscle (Fig. 4), coincident with the
upregulation of the expression of acidic fast-twitch skeletal muscle
TnT isoforms (26). There is no change in Tm isoform expression during
the development of the gastrocnemius, where only basic TnT isoforms are
expressed. It is interesting to speculate that the programmed,
exclusive expression of
-Tm in breast muscle may be a response to
the acidic TnT isoform expression, possibly also contributing to the
cooperativity of muscle activation seen in Fig. 6. Whether an
evolutionary adaptation is responsible for the coexpression of specific
TnT and Tm isoforms remains to be investigated.
Potential effect of the NH2-terminal
charge of TnT isoforms on muscle contractility.
Our data suggest that acidic and basic TnT isoforms may modulate the
Ca2+ sensitivity of muscle
contraction, implicating the NH2
terminus of TnT in this function. One possible mechanism for this
change in sensitivity may be dictated by the
NH2-terminal charge of TnT. By the
Gibbs-Donnan equilibrium, acidic TnT isoforms with negative charges at
the NH2 terminus may increase the
local free Ca2+ concentration at
the thin filament to a level higher than that in the bulk solution. In
effect, this increased local Ca2+
concentration would be available to bind to TnC and trigger
contraction. This would be relevant for both skinned and intact muscle
fiber preparations. In this case, detailed experiments elucidating the molecular distance between the TnT
NH2 terminus and the regulatory Ca2+-binding sites of TnC would
determine the magnitude of this effect.
The effect of TnT isoforms on the actomyosin system may also be
directly due to different interactions with the other components in the
thin-filament regulatory assembly (39), such as Tm dimers, which, in a
simple model, are believed to contribute to the steric block of
F-actin-active sites and prevent myosin head attachment (for recent
review see Ref. 38). The predominant structural difference among TnT
isoforms lies in the NH2 terminus,
a region that has been shown to interact with the head-to-tail overlap of Tm dimers (4). The NH2 and COOH
termini of
- and
-Tm show a conserved, high proportion of charged
amino acids (Asp, Glu, His, Lys, Arg) (21), and the potential of ionic
interactions between this region of Tm and the charged
NH2 terminus of TnT is a potential
mechanism through which TnT-Tm interactions may modulate the response
of the thin filament to Ca2+
activation. Unique interactions between TnT and Tm isoforms have been
shown in experiments by Pearlstone and Smillie (30), in which rabbit
fast-twitch skeletal muscle TnT fragments showed different binding
affinities to various tropomyosin isoforms. Furthermore, the variable
NH2 terminus of TnT isoforms may
dictate changes in the overall conformation and function of the protein (25, 41). More specifically, these differences in tertiary structure
among TnT isoforms may affect their interaction with TnI, TnC, and Tm,
providing another mechanism through which the regulated expression of
TnT isoforms may affect thin-filament activation. Altogether, TnT may
be a central molecule in modulating the function of the thin filament
through expression of multiple classes of isoforms.
 |
ACKNOWLEDGEMENTS |
We thank Jill Jin for the illustration of chicken muscle anatomy in
Fig. 3, Dr. Larry Smillie for the TnC expression plasmid, and Dr. Jim
Lin for the CH1 MAb.
 |
FOOTNOTES |
This study was supported in part by grants from the Medical Research
Council of Canada and the Heart and Stroke Foundation of Canada to
J.-P. Jin and National Institute of Arthritis and Musculoskeletal and
Skin Diseases Grant AR-42652 to H. Granzier.
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: J.-P. Jin, Dept.
of Physiology and Biophysics, Case Western Reserve University School of
Medicine, 10900 Euclid Ave., Cleveland, OH 44106-4970 (E-mail:
jxj12{at}po.cwru.edu).
Received 16 November 1998; accepted in final form 12 February
1999.
 |
REFERENCES |
1.
Akella, A. B.,
X. L. Ding,
R. Cheng,
and
J. Gulati.
Diminished Ca2+ sensitivity of skinned cardiac muscle contractility coincident with troponin T band shifts in diabetic rats.
Circ. Res.
76:
600-606,
1995[Abstract/Free Full Text].
2.
Anderson, P. A.,
A. Greig,
T. A. Mark,
N. N. Malouf,
A. E. Oakeley,
R. M. Ungerleider,
P. D. Allen,
and
B. K. Kay.
Molecular basis of human cardiac troponin T isoforms expressed in the developing, adult, and failing heart.
Circ. Res.
76:
681-686,
1995[Abstract/Free Full Text].
3.
Breitbart, R. E.,
and
B. Nadal-Ginard.
Complete nucleotide sequence of the fast skeletal troponin T gene: alternatively spliced exons exhibit unusual interspecies divergence.
J. Mol. Biol.
188:
313-324,
1986[Medline].
4.
Brisson, J. R.,
K. Golosinska,
L. B. Smillie,
and
B. D. Sykes.
Interaction of tropomyosin and troponin T: a proton nuclear magnetic resonance study.
Biochemistry
25:
4548-4555,
1986[Medline].
5.
Chase, P. B.,
and
M. J. Kushmerick.
Effects of pH on contraction of rabbit fast and slow skeletal muscle fibres.
Biophys. J.
53:
935-946,
1988[Abstract].
6.
Collins, J. H.
Myosin light chains and troponin C: structural and evolutionary relationships revealed by amino acid sequence comparisons.
J. Muscle Res. Cell Motil.
12:
3-25,
1991[Medline].
7.
Cooper, T. A.,
and
C. P. Ordahl.
A single cardiac troponin T gene generates embryonic and adult isoforms via developmentally regulated alternative splicing.
J. Biol. Chem.
260:
11140-11148,
1985[Abstract/Free Full Text].
8.
Fabiato, A.
Computer programs for calculating total from specified free or free from specified total ionic concentrations in aqueous solutions containing multiple metals and ligands.
Methods Enzymol.
157:
378-417,
1988[Medline].
9.
Fujita, S.,
K. Maeda,
and
Y. Maeda.
Expression in Escherichia coli and a functional study of a
-troponin T 25 kDa fragment of rabbit skeletal muscle.
J. Biochem. (Tokyo)
112:
306-308,
1992[Abstract].
10.
Godt, R. E.,
R. T. Fogaca,
and
T. M. Nosek.
Changes in force and calcium sensitivity in the developing avian heart.
Can. J. Physiol. Pharmacol.
69:
1692-1697,
1991[Medline].
11.
Granzier, H. L. M.,
and
T. Irving.
Passive tension in cardiac muscle: contribution of collagen, titin, microtubules, and intermediate filaments.
Biophys. J.
68:
1027-1044,
1995[Abstract].
12.
Granzier, H. L. M.,
J. Myers,
and
G. H. Pollack.
Stepwise shortening of muscle fibre segments.
J. Muscle Res. Cell Motil.
8:
242-251,
1987[Medline].
13.
Heeley, D. H.,
K. Golosinska,
and
L. B. Smillie.
The effects of troponin T fragments T1 and T2 on the binding of nonpolymerizable tropomyosin to F-actin in the presence and absence of troponin I and troponin C.
J. Biol. Chem.
262:
9971-9978,
1987[Abstract/Free Full Text].
14.
Jin, J.-P.,
A. Chen,
and
Q.-Q. Huang.
Three alternatively spliced mouse slow skeletal muscle troponin T isoforms: conserved primary structure and regulated expression during postnatal development.
Gene
214:
121-129,
1998[Medline].
15.
Jin, J.-P.,
Q.-Q. Huang,
H. I. Yeh,
and
J. J.-C. Lin.
Complete nucleotide sequence and structural organization of rat cardiac troponin T gene. A single gene generates embryonic and adult isoforms via developmentally regulated alternative splicing.
J. Mol. Biol.
227:
1269-1276,
1992[Medline].
16.
Jin, J.-P.,
and
J. J.-C. Lin.
Rapid purification of mammalian cardiac troponin T and its isoform switching in rat hearts during development.
J. Biol. Chem.
263:
7309-7315,
1988[Abstract/Free Full Text].
17.
Jin, J.-P.,
J. L.-C. Lin,
and
J. J.-C. Lin.
Troponin T isoform switching during heart development.
Ann. NY Acad. Sci.
588:
393-396,
1990.
18.
Jin, J.-P.,
and
L. B. Smillie.
An unusual metal-binding cluster found exclusively in the avian breast muscle troponin T of Galliformes and Craciformes.
FEBS Lett.
341:
135-140,
1994[Medline].
19.
Jin, J.-P.,
J. Wang,
and
J. Zhang.
Expression of cDNAs encoding mouse cardiac troponin T isoforms: characterization of a large sample of independent clones.
Gene
168:
217-221,
1996[Medline].
20.
Leavis, P. C.,
and
J. Gergely.
Thin filament proteins and thin filament-linked regulation of vertebrate muscle contraction.
CRC Crit. Rev. Biochem.
16:
235-305,
1984[Medline].
21.
Lees-Miller, J. P.,
and
D. M. Helfman.
The molecular basis for tropomyosin isoform diversity.
Bioessays
13:
429-437,
1991[Medline].
22.
Lin, J. J.-C.,
C. S. Chou,
and
J. L.-C. Lin.
Monoclonal antibodies against chicken tropomyosin isoforms: production, characterization, and application.
Hybridoma
3:
223-242,
1985.
23.
Martyn, D. A.,
and
A. M. Gordon.
Force and stiffness in glycerinated rabbit psoas fibres. Effects of calcium and elevated phosphate.
J. Gen. Physiol.
99:
795-816,
1992[Abstract].
24.
Nickel, R.,
A. Schummer,
and
E. Seiferle.
Anatomy of the Domestic Birds. Berlin: Verlag Paul Parey, 1977.
25.
Ogut, O.,
and
J.-P. Jin.
Expression, zinc-affinity purification and characterization of a novel metal-binding cluster in troponin T: metal-stabilized
-helical structure and effects of the NH2-terminal variable region on the conformation of intact troponin T and its association with tropomyosin.
Biochemistry
35:
16581-16590,
1996[Medline].
26.
Ogut, O.,
and
J.-P. Jin.
Developmentally regulated, alternative RNA splicing-generated pectoral muscle-specific troponin T isoforms and role of the NH2-terminal hypervariable region in the tolerance to acidosis.
J. Biol. Chem.
273:
27858-27866,
1998[Abstract/Free Full Text].
27.
Palmiter, K. A.,
Y. Kitada,
M. Muthuchamy,
D. F. Wieczorek,
and
R. J. Solaro.
Exchange of
- for
-tropomyosin in hearts of transgenic mice induces changes in thin filament response to Ca2+, strong cross-bridge binding, and protein phosphorylation.
J. Biol. Chem.
271:
11611-11614,
1996[Abstract/Free Full Text].
28.
Pan, B. S.,
A. M. Gordon,
and
J. D. Potter.
Deletion of the first 45 NH2-terminal residues of rabbit skeletal troponin T strengthens binding of troponin to immobilized tropomyosin.
J. Biol. Chem.
266:
12432-12438,
1991[Abstract/Free Full Text].
29.
Pearlstone, J. R.,
P. Johnson,
M. R. Carpenter,
and
L. B. Smillie.
Primary structure of rabbit skeletal muscle troponin-T. Sequence determination of the NH2-terminal fragment CB3 and the complete sequence of troponin-T.
J. Biol. Chem.
252:
983-989,
1977[Abstract].
30.
Pearlstone, J. R.,
and
L. B. Smillie.
Binding of troponin-T fragments to several types of tropomyosin.
J. Biol. Chem.
257:
10587-10592,
1982[Abstract].
31.
Perry, S. V.
Troponin T: genetics, properties and function.
J. Muscle Res. Cell Motil.
19:
575-602,
1998[Medline].
32.
Potter, J. D.
Preparation of troponin and its subunits.
Methods Enzymol.
85:
241-263,
1982[Medline].
33.
Reiser, P. J.,
M. L. Greaser,
and
R. L. Moss.
Developmental changes in troponin T isoform expression and tension production in chicken single skeletal muscle fibres.
J. Physiol. (Lond.)
449:
573-588,
1992[Abstract].
34.
Schachat, F. H.,
M. S. Diamond,
and
P. W. Brandt.
Effect of different troponin T-tropomyosin combinations on thin filament activation.
J. Mol. Biol.
198:
551-554,
1987[Medline].
35.
Schachat, F. H.,
J. M. Schmidt,
M. Maready,
and
M. M. Briggs.
Chicken perinatal troponin Ts are generated by a combination of novel and phylogenetically conserved alternative splicing pathways.
Dev. Biol.
171:
233-239,
1995[Medline].
36.
Smillie, L. B.,
K. Golosinska,
and
F. C. Reinach.
Sequences of complete cDNAs encoding four variants of chicken skeletal muscle troponin T.
J. Biol. Chem.
263:
18816-18820,
1988[Abstract/Free Full Text].
37.
Solaro, R. J.,
J. A. Lee,
J. C. Kentish,
and
D. G. Allen.
Effects of acidosis on ventricular muscle from adult and neonatal rats.
Circ. Res.
63:
779-787,
1988[Abstract].
38.
Squire, J. M.,
and
E. P. Morris.
A new look at thin filament regulation in vertebrate skeletal muscle.
FASEB J.
12:
761-771,
1998[Abstract/Free Full Text].
39.
Tobacman, L. S.
Structure-function studies of the amino-terminal region of troponin T.
J. Biol. Chem.
263:
2668-2672,
1988[Abstract/Free Full Text].
40.
Wang, J.,
and
J.-P. Jin.
Primary structure and developmental acidic to basic transition of 13 alternatively spliced mouse fast skeletal muscle troponin T isoforms.
Gene
193:
105-114,
1997[Medline].
41.
Wang, J.,
and
J.-P. Jin.
Conformational modulation of troponin T by configuration of the NH2-terminal variable region and functional effects.
Biochemistry
37:
14519-14528,
1998[Medline].
42.
Westfall, M. V.,
E. M. Rust,
and
J. M. Metzger.
Slow skeletal troponin I gene transfer, expression, and myofilament incorporation enhances adult cardiac myocyte contractile function.
Proc. Natl. Acad. Sci. USA
94:
5444-5449,
1997[Abstract/Free Full Text].
43.
Wu, Q.-L.,
P. K. Jha,
M. K. Raychowdbury,
Y. Du,
P. C. Leavis,
and
S. Sarkar.
Isolation and characterization of human fast skeletal troponin T cDNA: comparative sequence analysis of isoforms and insight into the evolution of members.
DNA Cell Biol.
13:
217-233,
1994[Medline].
44.
Yonemura, I.,
T. Watanabe,
M. Kirinoki,
J. Miyazaki,
and
T. Hirabayashi.
Cloning of chicken slow muscle troponin T and its sequence comparison with that of human.
Biochem. Biophys. Res. Commun.
226:
200-205,
1996[Medline].
Am J Physiol Cell Physiol 276(5):C1162-C1170
0002-9513/99 $5.00
Copyright © 1999 the American Physiological Society