From the
Abteilung Innere Medizin III (Kardiologie), Universität Heidelberg,
69115 Heidelberg, Germany, Institut für
Physiologie und Pathophysiologie, Medizinische Biophysik, Im Neuenheimer Feld
326, Universität Heidelberg, 69120 Heidelberg, Germany, and
||Institute of Biomedical Life Science, University
of Glasgow, G128QQ Glasgow, United Kingdom
Received for publication, April 1, 2003 , and in revised form, April 23, 2003.
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Although subsequent physiological studies in cardiac muscle recently identified S100A1 as a novel regulator of cardiac contractility being essential for cardiac reserve (811), significantly less is known regarding the impact of S100A1 on contractile properties of skeletal muscle. Biochemical and biophysical studies, however, have indicated that S100A1 can physically interact with the purified sarcoplasmic Ca2+ release channel/ryanodine receptor skeletal muscle isoform (RyR1)1 (12), the molecular entity that is the key substrate for sarcoplasmic reticulum (SR) Ca2+ release in striated muscle. However, it was unclear whether S100A1/RyR1 interaction is able to enhance Ca2+ release from the intact SR to increase contractile performance in skeletal muscle fibers. We therefore sought to investigate the role of S100A1 protein in saponin-skinned slow-twitch (Musculus soleus, Soleus) and fast-twitch (Musculus extensor digitorum longus, EDL) murine skeletal muscle fibers maintaining structural and functional integrity of the SR and the contractile apparatus. This approach enabled us to assess the effect of S100A1 on SR Ca2+ release and the interplay between preserved SR and contractile apparatus function (13).
In general, S100 proteins consist of an N-terminal part containing a non-conventional Ca2+-binding site of the EF-hand (helix-loop-helix) type and a C-terminal part containing a canonical EF-hand. The two parts are interconnected by an intermediate region, the "hinge region," and the C-terminal EF-hand is followed by a C-terminal extension. The hydrophobic hinge region and the C-terminal extension display the least amount of sequence identity among S100A1 members and are suggested to specify the biological activities of individual S100A1 proteins. S100 proteins typically display structural changes and exposure of hydrophobic surfaces upon Ca2+ binding, reminiscent of the sensor properties of calmodulin, to interact with their target proteins. Based on novel insights gathered from structural analysis of S100A1 protein (14), a S100A1 peptide model consisting of the region amino acids 216 (Fig. 1C, N-terminal (N)), amino acids 4254 (Hinge-region (H)), and amino acids 7594 (C-terminal (C)) devoid of Ca2+-binding motifs was synthesized to gain further insight into structure-function relationship of S100A1/RyR1 interaction as well as to exclude adverse Ca2+-buffering effects by the native protein (15, 16)
|
Taking advantage of chemically skinned skeletal muscle fibers, we were able to demonstrate for the first time that S100A1/RyR1 interaction can enhance SR Ca2+ release from the intact SR resulting in increased isometric force transients both in slow- and fast-twitch skeletal muscle. Importantly, S100A1 protein as well as the synthetic S100A1 peptide model (N/H/C) equally enhanced functional parameters, and these effects were found to be dose-dependent in a range of 0.00110 µM. Further testing of single S100A1 domains identified the hydrophobic C-terminal extension (aa 7594) as well as the hinge region (aa 4254) to differentially affect SR function. These effects are apparently based on enhanced SR Ca2+ release as S100A1 neither influenced SR Ca2+ uptake nor myofilament Ca2+ sensitivity/cooperativity in skeletal muscle fibers in our experimental setting. Thus, our data suggest a putative physiological role for S100A1 to serve as an endogenous enhancer of SR Ca2+ release in skeletal muscle.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Muscle Fiber Preparation and Experimental SolutionsAll of the animals were handled according to the guidelines of the animal care committee of the University of Heidelberg. Male BALB/c mice (36-months-old) were sacrificed by an overdose of carbon dioxide, and muscle fiber preparation was carried out as previously described (21, 22). Either EDL or Soleus was isolated, and a small fiber bundle containing two to four single fibers (between 80 and 150 µm in diameter and 34-mm-long) was dissected in paraffin oil. The fiber preparation was glued between a force transducer pin (AE801, Senso-Noras, Horton, Norway) and a micrometer-adjustable screw. All of the experiments were carried out at room temperature (2325 °C). All of the solutions were adjusted to pH 7.0. The free ion concentrations were calculated with the computer program REACT (version 2.0) from G. L. Smith (Glasgow, Scotland). Table I shows the concentrations of the solution used in the experiments. The high relaxation and the high activation solution contained 50 mM EGTA to buffer free Ca2+, whereas the low relaxing solution contained 0.5 mM EGTA and 49.5 mM 1,6-diamino hexane-N,N,N,N-tetraacetic acid (HDTA), which in contrast to EGTA has very low affinity to Ca2+. The skinning solution is obtained by the addition of 50 µg/ml saponin to the low relaxing solution. The release solution consisted of the low relaxing solution with 5 mM caffeine added. Loading solution contained 50 mM EGTA to clamp free Ca2+ to 0.4 µM (pCa 6.4). The solutions to measure the pCa-force relation were obtained by mixing high relaxing solution with appropriate amounts of high activating solution, and 5 mM caffeine added. All of the experiments were recorded using a strip chart recorder and were simultaneously digitally converted with an Axon Instruments Digidata 1200 board and interface (using the Axotape Software, version 2.0) and stored on the computer (22).
|
Assessment of Ca2+-induced Isometric Twitch Force and Ca2+ TransientsMuscle fibers were skinned for 5 min in skinning solution while the sarcomere length was adjusted to 2.6 ± 0.1 µm using the diffraction pattern of a helium-neon laser (22). Before loading the SR with the loading solution (pCa 6.4) for 1 min, the fibers were shortly immersed in release solution and high relaxing solution and then equilibrated for 2 min in low relaxing solution. Subsequently, the preparation was dipped for 1 s into the high relaxing solution and again for 2 min in low relaxing solution. The fibers were exposed to the release solution containing 5 mM caffeine until the initial force transient returned to the resting force level. Maximum force was measured in the high activating solution at pCa 4.28 and 5 mM caffeine. The fibers then were relaxed in high relaxing solution for 1 min to buffer Ca2+. Several control transients were recorded before the fiber was exposed to S100A1, and the experiment was repeated as outlined above. S100A1 protein or peptides were added to the low relaxing solution before and during release and to the high activating solution (22). The pCa-force relation in response to S100A1 interventions (0.00110 µM) was measured with six different Ca2+ concentrations (EDL, pCa 9.07, 5.91, 5.72, 5.49, 5.17, and 4.28; Soleus, pCa 9.07, 5.72, 5.49, 5.35, 5.17, and 4.28), each containing 5 mM caffeine. The EC50 and the Hill coefficient were obtained from a Hill-type fit (23). The EC50 value indicates the Ca2+ concentration needed for half-maximal isometric force activation, which is as a measure of Ca2+ sensitivity of the contractile apparatus. The Hill coefficient gives an indication of the maximum steepness of the sigmoidal curve. The correlation coefficients were calculated to determine the accuracy of the fit. The force transient was transformed into the corresponding free Ca2+ transient by using the individual pCa2+ force relation as a Ca2+ indicator and reversing each point of the force transients into the corresponding free Ca2+ level as previously described (22, 24, 25). Based on the fact that sensitivity of the Ca2+-regulatory proteins and the corresponding force development directly provide a measure of the free myofibrillar Ca2+, the pCa force relation relates free Ca2+ and force. Thus, the pCa-force relation can be used as a bioassay, which converts the rather slow force transients from the Ca2+ release from the SR into apparent Ca2+ transients (22, 24, 25).
Sarcoplasmic Ca2+ Uptake in EDL SR
VesiclesEDL muscles from hind legs of male Balb/c mice (36
months) were dissected and used for SR vesicle preparation as previously
published (26). SR vesicle
protein content was measured using the DMTM protein assay (Bio-Rad), and
aliquots were stored at 80 °C. Sarcoplasmic
Ca2+ uptake was measured as described elsewhere
(27). EDL SR vesicles (100
µg) were suspended in a 1.5-ml reaction solution (in mM: 120
KCl, 5 MgATP, 15 CrP, 1 MgCl, 25 HEPES, 20 K2Oxalate, 0.05
K2EGTA, pH 7.0) and equilibrated with 0.01 mM Fura-2
(Sigma) and 5 µM ruthenium while stirring in a cuvette (1.5 ml).
Ca2+ uptake measurements were started after the addition
of 67 µM CaCl2, resulting in an increase in free
[Ca2+] from 100 nM to 1µM.
The consequent decline of Fura-2 fluorescence ratio (340:380 nm) was a
reflection of SR Ca2+ uptake, and the fluorescence ratio
was recorded at 30 Hz using a spinning wheel spectrophotometer (Cairn
Research). The low pass filtered (3 db at 30 Hz) signal was digitized
and stored for later analysis. Ca2+ uptake rate
(dCa2+/dt; pmol Ca2+/s) for 100
µg of SR protein was obtained from the time constant (
) of
extrasarcoplasmic [Ca2+] decline.
was achieved
from best-fit single-exponential decay from experiments where free
[Ca2+]i in the cuvette is exceeding
1 µM. The relationship between given Ca2+
concentrations and the resulting fluorescence ratios was established with a
series of calibration experiments and analyzed according to Grynkiewicz et
al. (28).
Ca2+-binding constants were taken from Fabiato and
Fabiato (29) and Baudier
et al. (15,
16). For S100A1 interventions,
SR vesicles were preincubated with either 1 or 10 µM S100A1
protein or peptides (N/H/C) for 30 min while S100A1 storage buffer served as
control.
S100A1 ImmunoprecipitationsS100A1 protein levels in murine heart, EDL, and Soleus were assessed as previously described (30). Either untreated or saponin-skinned cardiac myocytes EDL and Soleus fibers were homogenized at 4 °C in 3 w/v phosphate-buffered saline with 5 mM EGTA/EDTA and protease inhibitor mixture (1836170, Complete Mini EDTA free, Roche Diagnostics) followed by centrifugation (10,000 x g for 15 min). The suspensions were rotated with bovine serum albumin-treated A/G-Sepharose for 30 min and centrifuged (800 x g) to remove proteins bound nonspecifically to A/G-Sepharose. The supernatants were then mixed with A/G-Sepharose and precipitating antibody for S100A1 (SA 5632) and incubated overnight at 4 °C. The samples were rotated for 30 min and centrifuged (800 x g), and pellets were washed three times with a buffer composed of 20 mM HEPES, pH 7.5, 150 mM NaCl, 1 mM EDTA, and 0.5% Tween 20. Samples were subjected to SDS-PAGE (420%), transferred to PVDF membrane, and probed with affinity-purified polyclonal S100A1-Ab (DAKO A5109). Blots were developed with the Avidix chemiluminescence detection system (Tropix, Applied Biosystems, Foster City, CA) and quantified by densitometry.
Statistical AnalysesData are presented as the mean ± S.E. Unpaired Student's t test and a two-way repeated ANOVA analysis were performed to test for differences between groups. A value of p < 0.05 was accepted as statistically significant.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Although it has been reported that even application of high concentrations
of the Ca2+-binding protein S100A1 protein (80
µM) to EGTA-buffered Ca2+ solutions caused
no change in free Ca2+ concentration
(31), we sought to introduce a
synthetic S100A1 peptide model devoid of Ca2+-binding
motifs both to exclude any artificial lowering of the free
Ca2+ concentration by S100A1 protein in our experimental
setting and to gain further insight into structure-function relationship of
S100A1 effects. Amino acid sequence alignment displayed in
Fig. 1C compares
S100A1 peptides defined by residues 216 (N-terminal
(N)), 4254 (Hinge-region (H)), and
7594 (C-terminal (C)) to the human S100A1 protein
primary sequence. The selected peptides encompass nearly 50% of the protein,
however, omitting both EF-hand Ca2+-binding domains.
Fig. 1D exhibits a
representative silver staining of the three S100A1 peptides resolved by
420% SDS-PAGE, visualizing the high purity that was confirmed by
analytical reverse-phase high performance liquid chromatography analysis (data
not shown). Molecular weight was determined by the use of matrix-assisted
laser desorption ionization-time of flight mass spectroscopy. Peak signals
yielded in the mass spectrum (N, 1653.26 ± 0.52 Da; H, 1376.17 ±
0.39 Da; C, 2257.10 ± 0.19; n = 5) were nearly identical to
the calculated peptide mass (N, 1652 Da; H, 1377 Da; and C, 2258 Da).
Immunoprecipitation of endogenous S100A1 protein levels in murine
cardiomyocyte, EDL, and Soleus muscle preparations visualized both by silver
staining (Fig. 1E,
upper panel) and Western blotting
(Fig. 1E, lower
panel) confirmed both differential S100A1 expression levels in murine
heart (16.87 ± 1.22 densitometric arbitrary units; 100%), Soleus (6.56
± 0.89 densitometric arbitrary units; 38% compared with heart), and EDL
(0.72 ± 0.04 densitometric arbitrary units; 4.26% compared with heart)
as well as S100A1 depletion of striated muscle following saponin treatment.
S100A1 monomer and dimer are indicated by single and double
asterisks, respectively.
S100A1 Increases Caffeine-induced SR Ca2+ Release and Ca2+-induced Isometric Force Transients in Murine EDL and Soleus Muscle FibersS100A1 has been reported to be mainly found in cardiac and slow-twitch skeletal muscle, whereas fast-twitch muscle fibers contain lower amounts of S100A1 protein (6, 32), which could be confirmed by S100A1 immunoprecipitation (Fig. 1E). Since S100A1 has been shown to interact with the SR Ca2+ release channel/ryanodine receptor (RyR1) reconstituted in lipid bilayers (12), we were interested whether S100A1/RyR1 interaction can modulate SR function and contractile properties of skeletal muscle fibers. To gain direct diffusional access for S100A1 protein and peptides to the myoplasm and its target proteins, the sarcolemma from small EDL and Soleus muscle bundles was rendered permeable by saponin treatment, thus leaving the SR and the contractile apparatus fully intact (24). Importantly, saponin treatment resulted in significant depletion of endogenous S100A1 protein levels both in EDL and Soleus muscle preparations (Fig. 1E). According to previous biophysical approaches (12), we first investigated SR Ca2+ efflux in response to acute application or preincubation (2 min) both of S100A1 protein and synthetic S100A1 peptides (N/H/C) in our experimental setting. Interestingly, in the presence of 0.5 mM free Mg2+ that is believed to inhibit channel opening by occupying the site for calcium activation of the RyR (33) S100A1 interventions failed to directly elicit SR Ca2+ release both in Soleus and EDL skeletal muscle preparations.
Therefore, we decided to investigate the impact of S100A1 on activated RyR-mediated SR Ca2+ release using caffeine. Caffeine was chosen to initiate RyR1 opening for its ability to increase Ca2+ sensitivity of the Ca2+ activation site on the SR Ca2+ release channel without appreciably affecting channel subconductance and sensitivity to endogenous regulators (e.g. ATP) (34, 35), which can be seen in the bell-shaped curve of channel activation versus [Ca2+] as a leftward shift of the ascending (activation) arm of the curve with little change in the descending (inhibition) limb (36).
Prior to S100A1 interventions, a series of caffeine-induced control
Ca2+ releases were established and EDL and Soleus fibers
were loaded with Ca2+ in such a manner that the peaks of
the Ca2+-induced force transients reached
2040% of the maximal isometric
Ca2+-dependent force. All of the force transients were
normalized to maximum force to correct for the rundown of the fiber.
Fig. 2 shows typical examples
for S100A1 protein (Fig. 2, A and
B) and the S100A1 peptide-(N/H/C)
(Fig. 2, E and
F) interventions on isometric force transients in EDL and
Soleus muscle fibers compared with control. The addition of S100A1 protein (1
µM) resulted in an equal reversible increase of the isometric
force transients both in EDL (+53.7 ± 10.1%, n = 6; *,
p < 0.01) and Soleus (+55.1 ± 4.12%, n = 5; *,
p < 0.01) compared with control
(Fig. 2I). Application
of the S100A1 peptide model (N/H/C) (1 µM) resulted in a similar
enhancement in peak amplitudes of the force transients in EDL (+59.9 ±
8.13%, n = 6; *, p < 0.01) and Soleus (+58.9 ±
9.31%, n = 7; *, p < 0.01) demonstrating identical
activity of the synthetic S100A1 model compared with the native protein
(Fig. 2I).
|
Corresponding Ca2+ transients for S100A1 protein and S100A1 peptide-(N/H/C) interventions in EDL and Soleus muscle fibers (Fig. 2, C and D and G and H) were obtained by transformation of the force transients using the inverse Hill function fitted to the individual pCa-force relationships as previously described (22) that were recorded for every fiber preparation (see "Experimental Procedures") (22, 24, 25). S100A1 protein and peptides (N/H/C) were found to equally enhance calculated peak Ca2+ values of the Ca2+ transients by +58.7 ± 5.13% (n = 6; *, p < 0.01) and +52.3 ± 11.3% (n = 6; *, p < 0.01) in EDL and by +45.9 ± 7.30% (n = 5; *, p < 0.01) and +48.3 ± 4.17% (n = 7; *, p < 0.01) in Soleus, respectively, compared with control Ca2+ transients (Fig. 2J). Moreover, integration of the area (time integral) of the calculated Ca2+ transient served as a relative indicator of the amount of Ca2+ released from the SR (21, 37). Normalized to the area under the control Ca2+ transients, S100A1 interventions (1 µM) significantly enhanced the caffeine-triggered amount of Ca2+ released by the SR in EDL (S100A1 protein: +98.0 ± 8.71%, n = 4; *, p < 0.01; S100A1 peptides (N/H/C): +91.0 ± 9.63%, n = 4; *, p < 0.01) and Soleus (S100A1 protein: +90.4 ± 12.3%, n = 5; *, p < 0.01; S100A1 peptides (N/H/C): +87.8 ± 12.4%, n = 5; *, p < 0.01) muscle fibers compared with control.
Given the equal bioactivity of S100A1 peptides (N/H/C) and S100A1 protein in both skeletal muscle isoforms, the testing of dose dependence was restricted to S100A1 peptides in EDL muscle preparations. Application of incremental concentrations of S100A1 peptides (N/H/C) in a range of 0.0011 µM revealed a dose-dependent enhancement of caffeine-induced normalized peaks of isometric force and Ca2+ transients, respectively (Fig. 3, A and B). However, increasing concentrations of S100A1 beyond 1 µM resulted again in diminished amplitudes of isometric twitch force and Ca2+ transients. With regard to three potential S100A1-binding domains that have been identified on each subunit of the RyR1 (12), these results support the notion of a biphasic Ca2+-dependent action of S100A1 as already described for calmodulin (35, 38).
|
As S100A1 has been shown to colocalize with the SR in skeletal muscle
(6) and inhibition of SERCA
activity accounts for increased peak force in skinned fiber preparations
(22), the effect both of
S100A1 protein and S100A1 peptides on SERCA activity was examined. SERCA
activity was assessed by oxalate-facilitated SR Ca2+
uptake measurements in SR vesicles from murine EDL. Ca2+
uptake was started by the addition of Ca2+ to a final
concentration >1 µM free [Ca2+], and
the decline of the Fura-2 fluorescence within the cuvette was recorded in the
presence of 5 mM ruthenium red to inhibit
Ca2+ release. Following calibration,
Ca2+ uptake rate (dCa2+/dt; pmol
Ca2+/s) for 1 µM free
[Ca2+] was calculated from the time constant of
[Ca2+] decline. Neither the addition of S100A1 protein
nor application of peptides (N/H/C) significantly altered SR
Ca2+ uptake rate compared with control (S100A1 protein:
1 µM, 359 pmol Ca2+/s; S100A1 protein: 10
µM, 367 pmol Ca2+/s; S100A1 peptide: 1
µM, 379 pmol Ca2+/s; S100A1 peptides: 10
µM, 375 pmol Ca2+/s; n = 6 for
each experiment; p = not significant versus control 354 pmol
Ca2+/s) (Fig.
3C). Moreover, S100A1 impact on SERCA activity and SR
Ca2+ uptake has also been tested in saponin-skinned
muscle fiber preparations. Fig.
3D shows that the addition of 1 µM S100A1
peptides to loading solution resulted in unchanged caffeine-induced isometric
peak force and corresponding SR Ca2+ release, confirming
that S100A1 did not alter SR Ca2+ loading in our
experimental setting. Similar results were obtained for 10 µM
S100A1 (data not shown).
S100A1 Domains Differentially Affect SR Function and Contractile Properties of Saponin-skinned Skeletal Muscle PreparationsBecause we could show that S100A1 protein enhances caffeine-induced SR Ca2+ release in chemically skinned skeletal muscle fibers, we next sought to gain further insight into the structure-function relationship of S100A1/RyR1 interaction. Introduction of the S100A1 peptide-(N/H/C) model enabled us to investigate differential biological effectiveness of distinct S100A1 domains. Fig. 4, BD, displays representative superimposed original tracings of caffeine-induced isometric force transients in EDL muscle preparations in response to S100A1-C, S100A1-H, or S100A1-N peptide interventions compared with control. The addition of the S100A1-C peptide (aa 7594) resulted in a nearly identical increase in the amplitude of the caffeine-evoked isometric force transient (+51.2 ± 12.1%, n = 5; *, p < 0.01 versus control; #, p < 0.03 versus S100A1-H peptide) compared with S100A1 protein or the combination of all of the three S100A1 peptides (N/H/C) (Fig. 4A). Further testing of the single S100A1-H peptide (aa 4254) revealed a minor but still significant enhancement of the amplitude of the caffeine-evoked isometric force transient (+18.5 ± 8.3%, n = 5, *, p < 0.05 versus control), whereas the addition of the S100A1-N peptide (aa 216) displayed no effect (Fig. 4A). Additive application both of the S100A1-C and S100A1-H peptide revealed no further increase above the single C-peptide, S100A1 peptides (N/H/C), or the native protein. Importantly, control experiments with degraded "scrambled" S100A1 peptides (N/H/C) that have been subjected to repeated freeze-thaw cycles and intense sonification did not influence the isometric force transient (Fig. 4A).
|
S100A1 Protein and Peptides Does Not Alter Myofilament Ca2+ Sensitivity in Murine Fast- and Slow-Twitch Muscle FibersS100A1 is assumed to bind to contractile filaments in striated muscle (5) and has been shown to modulate the function of sarcomeric proteins (8, 31, 39). Thus, it was necessary to determine whether S100A1 interventions modulate Ca2+ sensitivity of regulatory proteins (e.g. troponin C) of the contractile apparatus in skeletal muscle preparations in our experimental approach, which might have contributed to the observed modulation of peak twitch force. The pCa-force relationship was measured both at the beginning and the end of each protocol in EDL and Soleus skinned muscle fibers to determine both Ca2+ concentration for half-maximal isometric force activation as a measure of myofilament Ca2+ sensitivity (EC50 [Ca2+]) and the steepness of the sigmoidal curve (Hill coefficient) as a value for the Ca2+-dependent cooperative interactions among contractile and regulatory proteins. In addition, maximal tension development was recognized as a measure of Ca2+-dependent regulation of strong cross-bridge attachment between the thin and thick filament. In the presence of caffeine application of 1 µM S100A1 protein and S100A1 peptides (N/H/C), neither affected Ca2+ sensitivity nor cooperativity in Soleus (S100A1 peptides (N/H/C) (S100A1 peptides, pCa EC50 5.95 ± 0.03, nHill 2.5 ± 0.18, n = 4; S100A1 protein, pCa EC50 5.90 ± 0.02, nHill 2.35 ± 0.22, n = 4; control, pCa EC50 5.86 ± 0.05, nHill 2.47 ± 0.34, n = 4; p = not significant versus S100A1) and EDL (S100A1 peptides, pCa EC50 5.31 ± 0.07, nHill 4.41 ± 0.53, n = 4; S100A1 protein, pCa EC50 5.36 ± 0.10, nHill 4.22 ± 0.21, n = 4; control pCa EC50 5.33 ± 0.11, nHill 4.38 ± 0.31, n = 4, p = not significant versus S100A1) compared with control in our experimental setting. Accordingly, we found that S100A1 did not alter maximal Ca2+-dependent tension development with regard to normalized maximal tension development of control fibers. In addition, pCa-force relationship for each single S100A1 peptide intervention was also found to be unaltered (data not shown).
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
S100A1, a member of the Ca2+-binding protein family known as S100, is the most abundant S100 protein isoform in striated muscle (7, 41) and has been shown to colocalize, in particular, with the sarcoplasmic reticulum and the contractile apparatus (57, 32, 42). Although recent studies in vitro and in vivo have reported on S100A1 to play a crucial role in the regulation of cardiac contractility (811), the impact of S100A1 on skeletal muscle contractility remained elusive so far. Therefore, we took advantage of a specific membrane permeabilization with saponin maintaining the cellular architecture of the SR and contractile apparatus and controlling the intracellular milieu (43) to investigate the role of S100A1 in the regulation of SR Ca2+ efflux and contractile performance in skeletal muscle fibers.
In this study, we were able to demonstrate for the first time that
S100A1/RyR1 interaction results in increased contractile performance both of
slow- and fast-twitch skeletal muscle fibers because of enhanced SR
Ca2+ release. Importantly, despite different endogenous
S100A1 protein levels in skeletal muscle isoforms that have been estimated to
110 µM in slow- and 520 times less in
fast-twitch skeletal muscle (5,
6,
32), reconstitution of
chemically S100A1-depleted skeletal muscle fibers with either S100A1 protein
or S100A1 peptides (N/H/C) near to their native levels yielded similar effects
on isometric force transients and SR Ca2+ release.
Although slow- and fast-twitch skeletal muscles differ in many ways
(e.g. metabolism, protein isoform composition, and so forth), they
mainly express the same RyR isoform (RyR1), which could partially explain
identical effects of S100A1 on SR Ca2+ efflux in Soleus
and EDL muscle fibers.
However, in our experimental setting S100A1-mediated enhancement of SR Ca2+ release only occurred with caffeine while in its absence S100A1 interventions failed to activate RyR1 opening. Thus, at first glance our data appears to disagree with Treves et al. (12) who previously reported on S100A1 protein to directly increase open probability of the purified RyR1 reconstituted in lipid bilayers. One important reason for this apparent discrepancy may be that these experiments were carried out in the absence of Mg2+ (12), whereas in contrast, S100A1 effects on SR Ca2+ release in saponin-skinned skeletal muscle preparations were studied in the presence of Mg2+ near its native concentrations. This is essential to note because Mg2+ is as a central inhibitor of Ca2+-dependent activation of the release channel and its physiological concentration near 1 mM is necessary in maintaining the RyR1 channels closed at rest (33, 44, 45). S100A1 effects on single channel-gating properties therefore occurred under experimental conditions when the channel was strongly sensitized to Ca2+-dependent activation. Because the mechanism of caffeine is also based on increased channel sensitivity to activation by Ca2+ (35, 36, 46) both studies consistently show that S100A1 protein principally enhances RyR1 opening under conditions that sensitize the channel to Ca2+ (absence of Mg2+, caffeine).
Taken together, S100A1 appears to directly activate RyR1 opening in the absence of Mg2+ while physiological levels of Mg2+ effectively prevent this effect. Therefore, we propose that under physiological conditions that the L-type Ca2+ channel voltage-gated control mechanism and the presence of Mg2+ provide the intrinsic mechanisms to avoid spontaneous and/or sustained RyR1 opening via S100A1. However, once RyR1 activation is promoted, e.g. by drug- or voltage-induced opening of the channel, we speculate that S100A1 protein enhances activated SR Ca2+ release by increasing the channel open probability. In addition, it seems noteworthy that dose-dependent S100A1-mediated amplification of SR Ca2+ release in our experimental setting first occurred at nanomolar S100A1 concentrations similar to effective S100A1 concentrations reported by Treves et al. (12). This finding strongly supports the notion that even low native S100A1 protein levels as found in EDL muscle are already sufficient to regulate SR function and contractile performance. Thus, taking advantage of a more physiological approach, we speculate that S100A1 might rather serve as an endogenous enhancer of SR Ca2+ release in skeletal muscle than to directly open the SR Ca2+ release channel.
Based on primary sequence alignment (47) and three-dimensional reconstruction of S100A1 protein explored by NMR spectroscopy (14), we next applied a synthetic S100A1 peptide model to gain further insight into structure-function relationship of S100A1/RyR1 interaction. Importantly, as described above application for all of the three S100A1 peptides (N/H/C), omitting both Ca2+-binding loops revealed similar effects in slow- and fast-twitch skeletal muscle preparations compared with the native protein. Testing of single S100A1 peptides revealed that at least the S100A1 C-terminal amino acid sequence 7594 exerts nearly an identical biological activity as the native protein. Moreover, the S100A1 hinge region encompassing the amino acid sequence 4254 also displayed biological activity albeit less than the S100A1 C terminus, whereas the N-terminal extension displayed no effect on SR Ca2+ release.
Thus, our data support previous assumptions that both the C-terminal
residue and the hinge region, which are buried in the apoform
(48) and exposed in the
calcium-bound form (49),
mediate selectivity in S100 protein target binding and biological activity
while the N-terminal extension is recognized to stabilize the dimeric
structure of S100 proteins
(50). In addition, we were
able to show that different S100A1 domains exert differential biological
activity because the C-terminal extension was found to be 23 times
more effective than the linker region. These findings seem to be specific for
S100A1 interventions as the characterization of protein and peptide
preparations revealed highest purity and integrity of the compounds, whereas
denatured S100A1 peptides did not yield any biological effect.
Because myofilament Ca2+ sensitivity is another important factor that essentially contributes to the regulation of contractile force in skeletal muscle (43), we analyzed the impact of S100A1 interventions on the pCa-force relationship. Although an altered Ca2+ affinity of myofilament-associated regulator proteins by S100A1 could have contributed to the observed increase in peak force, S100A1 neither influenced Ca2+ sensitivity nor cooperativity of the contractile apparatus in our experimental setting. Further, Adhikari and Wang (31) showed that S100A1 protein can decrease myofilament Ca2+ sensitivity of skinned rabbit psoas skeletal muscle fibers. However, it should be noted that the study by Adhikari and Wang (31) was carried out at a shorter sarcomere length (2.12.2 µm) and that there are major differences with respect to muscle-specific and species-specific fiber-type composition (5153) to explain the different findings of S100A1 effects on pCa-force relationship.
Because previous studies reported a possible interaction of S100A1 with the skeletal muscle SR Ca2+-ATPase isoform (6), we sought to investigate whether S100A1 might affect SR Ca2+ uptake in skeletal muscle by the use of fluorescence-based Ca2+ uptake measurements in purified SR vesicles. Consistent with reports by Fano et al. (54), neither S100A1 protein nor S100A1 peptides (N/H/C) were found to alter SR Ca2+ uptake rate in our assay system. Because purification of SR vesicles may somewhat alter the responsiveness to endogenous regulators because of the loss of membrane compounds, we additionally assessed the effect of S100A1 on SR Ca2+ loading in saponin-skinned fiber preparations. In accordance with our fluorescence-based measurements, SR Ca2+ load as determined by caffeine-induced contractures was similar for S100A1 interventions and controls in skeletal muscle fibers. Thus, in skeletal muscle, S100A1 does not appear to indirectly modulate Ca2+-activated force by changing the rate of SR Ca2+ uptake.
In conclusion, we have shown the ability of S100A1/RyR1 interaction to enhance Ca2+ efflux from the intact SR in saponin-skinned skeletal muscle fibers resulting in increased contractile performance. Introducing a novel synthetic S100A1 peptide model allowed us to identify specific S100A1 domains that are critically implicated in the structure-function relationship of S100A1/RyR1 interaction. Interestingly, despite higher endogenous S100A1 protein levels in slow- than in fast-twitch skeletal muscle, both skeletal muscle isoforms exerted identical responsiveness to S100A1 interventions. Given that S100A1 has recently been proven to play a central role in the regulation of myocardial contractile performance (811), we propose an important physiological role for S100A1 in skeletal muscle in vivo to serve as an endogenous enhancer of activated RyR1-mediated sarcoplasmic Ca2+ release, thereby enhancing skeletal muscle contractile performance. However, further experiments taking advantage of in vivo genetic manipulation of endogenous S100A1 protein levels in skeletal muscle will have to finally address this issue.
![]() |
FOOTNOTES |
---|
These authors contributed equally to this work.
¶ Supported by the Boehringer Ingelheim Stiftung.
** These authors contributed and supported this work equally.
To whom correspondence should be addressed: Institut für Physiologie und
Pathophysiologie, Im Neuenheimer Feld 326, Universität Heidelberg, 69120
Heidelberg, Germany. Tel.: 49-6221-54-4065; Fax: 49-6221-54-4123; E-mail:
rainer.fink{at}agfink.de.
1 The abbreviations used are: RyR, release channel/ryanodine receptor;
skeletal muscle isoform; SR, sarcoplasmic reticulum; Soleus, M.
soleus; EDL, M. extensor digitorum longus; aa, amino acid;
ANOVA, analysis of variance; N, N-terminal; H, hinge region; C, C-terminal;
pCa, log[Ca2+](M); SERCA, SR
Ca2+-ATPase.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|