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Calcium-dependent movement of troponin I between troponin C and actin as revealed by spin-labeling EPR
BBRC Biochemical and Biophysical Research Communications 340 (2006) 462–468 www.elsevier.com/locate/ybbrc
Calcium-dependent movement of troponin I between troponin C and actin as revealed by spin-labeling EPR Tomoki Aihara, Shoji Ueki, Motoyoshi Nakamura, Toshiaki Arata
*
Department of Biological Sciences, Graduate School of Science, Osaka University and CREST/JST, Toyonaka, Osaka 560-0043, Japan Received 17 November 2005 Available online 15 December 2005
Abstract We measured EPR spectra from a spin label on the Cys133 residue of troponin I (TnI) to identify Ca2+-induced structural states, based on sensitivity of spin-label mobility to flexibility and tertiary contact of a polypeptide. Spectrum from Tn complexes in the Ca2+ state showed that Cys133 was located at a flexible polypeptide segment (rotational correlation time s = 1.9 ns) that was free from TnC. Spectra of both Tn complexes alone and those reconstituted into the thin filaments in the +Ca2+ state showed that Cys133 existed on a stable segment (s = 4.8 ns) held by TnC. Spectra of reconstituted thin filaments (Ca2+ state) revealed that slow mobility (s = 45 ns) was due to tertiary contact of Cys133 with actin, because the same slow mobility was found for TnI–actin and TnI–tropomyosin–actin filaments lacking TnC, T or tropomyosin. We propose that the Cys133 region dissociates from TnC and attaches to the actin surface on the thin filaments, causing muscle relaxation at low Ca2+ concentrations. 2005 Elsevier Inc. All rights reserved. Keywords: Muscle regulation; Actin; Troponin; Tropomyosin; Spin labeling; EPR
Contraction of vertebrate striated muscles is regulated by troponin (Tn)/tropomyosin complexes located on the actin thin filaments [1–6]. Troponin is a calcium binding protein, and its sensitivity to calcium concentration is important for its regulation. Tn is composed of three heterotrimeric subunits, i.e., a calcium (Ca2+) binding subunit (TnC), an inhibitory subunit (TnI), and a tropomyosinbinding subunit (TnT). The crystal structure of TnC suggested that the mechanism underlying its activation involved Ca2+ binding to two regulatory sites, inducing movement of helices in the N-terminal domain, and so exposing a hydrophobic patch and further augmenting interaction with TnI [7]. This mechanism was consistent with that described in other structural studies [8–11]. TnI contains a few functional regions at the C-terminus. Its inhibitory segment (amino acids 104–115) and second actin-binding site (amino acids 140–148) can bind to actin and inhibit actomyosin ATPase activity [12–17]. Moreover, *
Corresponding author. Fax: +81 6 6850 5441. E-mail address: [email protected] (T. Arata).
0006-291X/$ - see front matter 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2005.12.030
the switch segment (amino acids 116–131) between two actin-binding sites is thought to interact with the N-lobe of TnC in the presence of Ca2+ [18] and to dissociate from it by binding the second site to actin in the absence of Ca2+. Using fluorescence resonance energy transfer (FRET), a number of studies showed that Cys133 near the TnI switch segment underwent Ca2+-dependent movement around TnC and actin subunits on the thin filaments [19–22]. Crystal structures of cardiac and skeletal TnC/TnI/TnT complexes have been solved, and showed that the switch segment of TnI was clutched by the N-lobe of TnC in the +Ca2+ state [23,24], but was disordered in the Ca2+ state [24]. Therefore, it is important to understand how Tn complexes undergo Ca2+-triggered structural changes on the thin filaments and in muscle fibers. Site-directed spin-labeling electron paramagnetic resonance (SDSL-EPR) spectroscopy has been used to determine secondary, tertiary, and quaternary structures, and associated conformational changes of proteins [25,26]. Depending on the rigidity of the attached spin label, it is possible to investigate steric restrictions imposed by the environment
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of the label, motion of a segment of a polypeptide chain, or Brownian diffusion of a whole molecule [27]. Onishi et al. [28] reported that a spin label, presumably positioned on Cys133 in TnI, showed Ca2+ -dependent immobilization in Tn complexes. Here, we confirmed that removal of Ca2+ increased the mobility of the side-chain spin label on Cys133, located near the switch segment of the rabbit TnI complexes in solution. According to the mobility–structure relationships in standard proteins [27], detection of a mobile spectrum arises from a loop-like flexible structure, while that of an immobile spectrum arises from a spin label on a stable loop or a helix surface. With the reconstituted thin filaments, Ca2+ removal resulted in opposite effects, i.e., it induced rigid immobilization of the spin labels, whereas the spectrum in the presence of Ca2+ was the same as that in Tn complexes. The rigidly immobilized component of spin labels was also observed in reconstituted TnI–actin and TnI–tropomyosin–actin filaments. This suggests that the side chain of Cys133 makes tertiary contact with the actin surface after forming a flexible structure free from TnC, upon Ca2+ removal. Therefore, we propose that the switch segment undergoes a Ca2+-dependent movement between TnC and actin on the thin filaments. Materials and methods Reagents. 4-Maleimido-2,2,6,6-tetramethyl-1-piperidinoxy (MSL) was from Sigma Chemicals. The BCA–protein assay reagents used to measure protein concentrations were from Pierce Chemicals. Other reagents were of analytical grade. Protein preparation. Rabbit muscle protein was prepared from back and leg muscles, and chicken from breast muscles. Actin was prepared from acetone powder of rabbit skeletal muscles, according to the method of Spudich and Watt [29]. Native troponin complexes (TnC–I–T complexes) and tropomyosin were prepared from rabbit and chicken skeletal muscle residues after extraction of myosin by the method of Ebashi et al. [30]. After dialysis with 20 mM Tris–HCl (pH 7.5), 30 mM KCl, 0.1 mM CaCl2 (buffer A), and 1 mM DTT, troponin was purified using a Q-Sepharose Fast Flow column (2.5 · 10 cm) and eluted with a linear 30–400 mM KCl gradient. Fractions containing TnC–I–T complexes were identified, collected by measuring absorption at 280 nm and by SDS– PAGE, and finally dialyzed with buffer A containing 1 mM DTT. Spin labeling of troponin complexes. Prior to spin labeling, purified troponin complexes were passed through a Sephadex G-25 column (1 · 20 cm) equilibrated with buffer A without DTT. Then, troponin (10– 40 lM) was reacted with an equimolar amount of MSL overnight in the dark at 4 C. The following day, unreacted MSL was removed by passing labeled troponin (MSL-Tn) through a Sephadex G-25 column equilibrated with buffer A. When removing DTT or unreacted MSL for large-scale preparations (>20 ml), dialysis using buffer A was done instead of using a Sephadex G-25 column. For EPR measurements, the MSL-labeled troponin solution was concentrated up to 1–3 mg/ml (10–40 lM) and examined after 0.1 mM CaCl2 or 1 mM EGTA was added in buffer A to achieve the +Ca2+ state or Ca2+ state, respectively. MSL-labeled TnI (MSL–TnI) was isolated from MSL–Tn complexes by the method of Ojima and Nishita [32]. MSL–Tn was dialyzed against 6 M urea, 30 mM KCl, 1 mM EDTA, and 10 mM Tris–HCl (pH 7.6), and applied to a CM-Toyopearl 650 M column (2.5 · 10 cm). Following preequilibration with the same buffer (to wash out isolated TnC), MSL–TnI was eluted with a linear gradient of 30–300 mM KCl, before eluting TnT. Fractions containing MSL–TnI were identified by measuring absorption at 280 nm and by SDS–PAGE. Eluted MSL–TnI was gently dialyzed against 100 mM KCl and 20 mM Tris–HCl (pH 7.5). After insoluble
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components were removed by centrifugation, the MSL–TnI preparation was used for reconstitution of thin (TnC–I–T-tropomyosin–actin) Tn-actin and TnI–tropomyosin–actin filaments. Reconstitution of thin filaments G-actin was incubated in 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, and 4 mM MgCl2 for 30 min. Tropomyosin and MSL–Tn were added to G-actin solution (ca. 100 lM, 0.3–0.5 ml), in the presence of 1 mM CaCl2 (+Ca2+ state) or 5 mM EGTA (Ca2+ state), and further incubated for 1 h. Molar ratio was set as 1:1:3 for tropomyosin, MSL–Tn complexes, and actin. Then, the mixture was centrifuged at 100,000g for 30 min, and the pellet co-sedimented with MSL–Tn was finally resuspended in a small volume (20–100 ll) of 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, 4 mM MgCl2, and 1 mM CaCl2 or 5 mM EGTA. Amounts of TnC, TnI, TnT, tropomyosin, and actin were assessed by SDS–PAGE. The suspension, containing 20–40 lM MSL–troponin, was used for EPR measurements. TnI–actin and TnI–tropomyosin–actin were also prepared as described above. After G-actin (ca. 100 lM, 0.3–0.5 ml) was incubated in 100 mM KCl and 4 mM MgCl2 for 30 min, tropomyosin was mixed with actin solution at a molar ratio of 1:3 with G-actin. Isolated MSL–TnI was mixed with actin or actin–tropomyosin solution at a molar ratio of 1:3 with G-actin and incubated for 1 h. The mixture was centrifuged at 100,000g for 30 min, and the pellet co-sedimented with MSL–TnI was finally resuspended in a small volume (ca. 20 ll) of 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, and 4 mM MgCl2. Amounts of TnI, tropomyosin, and actin were assessed by SDS–PAGE. The suspension, containing 20– 40 lM MSL–TnI, was used for EPR measurements. ATPase assay. Inhibitory activity of troponin was measured using an actomyosin ATPase assay. Troponin (1–4 lM) was mixed with 1 lM tropomyosin, 3.5 lM actin, and 0.6 lM myosin in 0.1 M KCl, 4 mM MgCl2, 1 mM CaCl2, and 20 mM Tris–HCl (pH 7.5). The ATPase reaction was coupled with a regeneration system as described by Matsuo et al. [33], where 2 mM phosphoenolpyruvic acid, 0.3 mM NADH, 38 U/ ml pyruvate kinase, and 50 U/ml lactate dehydrogenase were mixed. The reaction was started at 25 C by addition of 100 lM ATP, and time course of NADH absorption at 340 nm was followed for 10 min. Next, 5–10 mM EGTA was mixed with the sample and followed for 10 min. Velocities of the decrease in absorption were compared on charts according to the methods of Nakamura et al. [11] and were defined as: [ATPase (+Ca2+) activity ATPase(Ca2+) activity]/ATPase (+Ca2+) activity. EPR measurements and analysis EPR spectroscopy was performed using a Bruker ELEXSYS E500 spectrometer equipped with a dielectric resonator. Sample solutions (ca. 15 ll) were loaded into capillaries (inside diameter, 1.0 mm), inserted into the resonator, and EPR spectra acquired using 1-G field modulation amplitude at 100 kHz and 5-mW incident microwave power at 25 C. The rotational correlation time (s) was determined for the original spectrum and for those of fast or slow components produced by computer subtraction. The rotational correlation time in the fast motion regime (s 109 s) was calculated according to the equation of Redfield [34]: s = 1.19 · 109 DH0{(V0/V+1)0.5 + (V0/V1)0.52}, where DH0 is the peak-to-peak line width of the central line and Vm is the peak-to-peak height [34]. The rotational correlation time in the slow motion regime (s 109 2 · 107 s) was calculated according to an equation from Freed and co-workers [35]: s = a (1Teff/Tmax)b, where a = 5.4 · 1010 s, b = 1.36, Tmax (=35 G) is the rigid limit for a particular spin, and 2Teff is the width between the low-field and high-field absorption peaks [35]. Further, we plotted the reciprocal second moment ÆHæ2 (G2) versus the reciprocal central line width ÆDH0æ calculated from EPR spectra, according to Mchaourab et al. [27]. The following equation was used: P P hHi2 ¼ Ai DB2i = Ai , where Ai is the intensity at i field on the absorption spectrum and DBi (G) is the absolute value at i field assessed
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from the peak of the central extreme. We also characterized the secondary structure where a spin label existed.
Results and discussion Spin labeling of Cys133 on TnI We used SDSL-EPR spectroscopy to determine conformational changes in the C-terminal region of TnI in troponin complexes. Rabbit skeletal TnI was exclusively labeled with MSL because Cys133 near the switch segment of TnI was the most reactive cysteine in the complex state as reported by Chong and Hodges [31]. We investigated the effects of spin labeling. Actomyosin ATPase of MSL-Tn inhibited 85–95% of troponin labeling, indicating that spin labeling did not significantly affect inhibition by actomyosin ATPase. Labeling efficiency for rabbit troponin, estimated from double integration of the spectrum, was more than 0.8 mol/mol troponin complexes. For chicken troponin where Cys133 was substituted with another amino acid residue, efficiency was about 0.5–0.6 mol/mol troponin complexes. The lower reactivity for the other cysteines of chicken troponin suggested that for rabbits the fraction of spin-labeled cysteines other than Cys133 was very small [31,36]. Mobility of the spin label attached to Cys133 of TnI in troponin C–I–T complexes We measured EPR spectra from the spin label (MSL) of Cys133 of TnI in ternary TnC–I–T complexes to investigate conformational changes of the switch segment in
TnI, induced by Ca2+. Fig. 1 (left) shows spectra of rabbit skeletal troponin complexes in the presence and absence of Ca2+. In the +Ca2+ state (top), absorption line shapes were broad, and the two extremes at the low- and high-field were small relative to the central peak. From the spectral parameters DH0 = 3.0–3.4, V0/V1 = 7.0–13.0, and V0/V+1 = 2.1–2.4 (see Materials and methods), we deduced a slow motion of rotational correlation time s = 4.80 ± 1.57 ns (Table 1). In the Ca2+ state (1 mM EGTA), sharp peaks (at ca. 3420 G) appeared with a very small broad peak (at ca. 3410 G). Similar changes were seen at high magnetic field (at ca. 3450 G). Sharp major peaks (DH0 = 2.4–2.6, V0/V1 = 3.9–6.2, and V0/V+1 = 1.3–1.6) corresponded to a correlation time of s = 1.93 ± 0.57 ns (Table 1). These spectral changes were almost the same as those reported by Onishi et al. [28]. Excess Ca2+ (final concentration, 10 mM) was added to achieve the +Ca2+ state, and the spectrum returned to the original one, indicating a reversible reaction. Similar results were obtained for binary complexes of MSL-labeled TnI and TnC (Table 1). Chicken skeletal troponin was also prepared and measured under the same conditions. In chicken TnI, asparagine was substituted for Cys133 near the switch segment. Fig. 1 shows spectra of rabbit troponin (left) compared with those of chicken (right). Chicken spectra showed weak signal intensity compared with those of rabbit. Broad peaks (2Teff = ca. 63.0 G) at ca. 3410 and ca. 3470 G indicated rigid immobilization (s = 12–13 ns). Similar but very small peaks (at ca. 3410 and ca. 3470 G) in the slow motion regime were seen from rabbit troponin in the Ca2+ state but not in the +Ca2+ state. Spectra were not sensitive to Ca2+ concentration. Therefore, most of the spin labels were
Fig. 1. EPR spectra of spin-labeled troponin complexes from rabbit (left) and chicken (right) skeletal muscles in the +Ca2+ (top) and Ca2+ (bottom) states.
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468 Table 1 Summary of the rotational correlation time (s) of the spin label in TnI of rabbit s (ns)
TnC–I TnC–I–T TnC–I–T–tropomyosin–actin TnI–actin TnI–tropomyosin–actin
Ca2+ +Ca2+ Ca2+ +Ca2+ Ca2+ +Ca2+
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Cys64 of TnI; and Cys98 of TnC) did not result in mobility changes of spin labels attached on the thin filaments. MSL–TnI reconstituted with actin and tropomyosin–actin
Fast
Slow
1.67 3.26 1.93 ± 0.57a 4.80 ± 1.57a 4.76c (72%)b 4.76 2.61c (55%) 2.61c (33%)
— — — — 45.2c (28%) — 45.2c (45%) 43.0c (67%)
a
Values are means ± standard deviation for three preparations. Values in parentheses are the populations of the fast and slow components. c Rotational correlation times were determined for the spectra produced by spectral subtraction. The slow component of TnC–I–T–tropomyosin– actin (Ca2+) was obtained by digital subtraction of the spectrum of TnC–I–T–tropomyosin–actin (+Ca2+) containing 72% of the total spins from the whole spectrum. The fast component of TnI–actin was produced by subtracting 45% of the total spins of the slow component of TnC–I–T– tropomyosin–actin (Ca2+) from the whole spectrum. The slow component of TnI–tropomyosin–actin was produced by subtracting the fast component of TnI–actin containing 33% of the total spins from the whole spectrum. b
used to selectively label Cys133 of rabbit TnI and showed mobility changes induced by Ca2+ while spin labels of any other cysteines (presumably Cys48 and Cys64 of TnI; and Cys 98 of TnC) did not result in any spectral changes. MSL–TnI in reconstituted thin filaments To examine how spin-label mobility of Cys133 in TnI changes in response to Ca2+ on thin filaments, we reconstituted them with MSL–Tn–tropomyosin, and actin. Fig. 2 shows EPR spectra from rabbit skeletal thin filaments in the presence (A) and absence (B) of Ca2+. In the +Ca2+ state, spectrum shape mimicked that of troponin C–I–T complexes alone. The rotational correlation time was 4.72 ns (Table 1). On the other hand, in the Ca2+ state, a marked shoulder appeared at the lower field (about 3410 G). Similar corresponding changes were seen at a higher magnetic field (3450 G). The shoulder indicates a component of slow motion. The rotational correlation time was calculated as 45.2 ns from 2Teff (67.3 G) of the spectrum produced by spectral subtraction of the spectrum of +Ca2+ state containing 72% of the total spins from that of the Ca2+ state. We could not observe any enhancement of the fast motional component by adding EGTA, unlike MSL–Tn alone. These results were very similar to those observed for MSL–TnI reconstituted into myofibrils [37]. The experiments with the thin filaments were also performed using chicken troponin complexes. Spin-labeled chicken TnC–I–T complexes and rabbit actin were mixed and reconstituted into thin filaments. The resulting spectrum was the same as that of TnC–I–T complexes alone and revealed no Ca2+ effects (data not shown). This indicated that cysteines other than Cys133 (presumably Cys48 and
To examine whether Ca2+-removal induced immobilization of the spin label on Cys133 of TnI in thin filaments, and that it was due to the binding of the side chain of Cys133 to actin or TnT/C, we reconstituted isolated MSL–TnI with actin with or without tropomyosin. Figs. 2C and D show spectra from TnI–actin and TnI–tropomyosin–actin, respectively. No Ca2+ effect was observed, unlike in the thin filaments (TnC–I–T–tropomyosin–actin). Both spectra, especially from TnI–tropomyosin–actin, were very similar to that from thin filaments reconstituted in the Ca2+ state. Both spectra consisted of two distinct components, i.e., fast and slow motions. The slow component (2Teff = 67.3 G) was close to the motion observed in thin filaments in the Ca2+ state (s = 45 ns). Therefore, the fast component of TnI–actin was easily produced by subtracting 45% of the total spins of the slow component obtained by decomposition of the two spectra of TnC–I– T–tropomyosin–actin shown in Figs. 2A and B. From the spectral parameters DH0 = 3.1, V0/V1 = 4.5, and V0/ V+1 = 1.7, we deduced rotational correlation time s = 2.6 ns for the fast component (Table 1). This fast component containing 33% of the total spins was subtracted from the spectrum of TnI–tropomyosin–actin (Fig. 2D) to produce the same slow component again (2Teff = 67.2 G, s = 43 ns). These results were a strong indication that Cys133 in TnI interacted directly with actin in thin filaments in the Ca2+ state, because it could interact with actin in the same way, irrespective of troponin or tropomyosin. Secondary structure prediction from side-chain mobility of the spin label Electron paramagnetic resonance (EPR) spectroscopy allows the investigation of motional sensitivity within a nanosecond to submillisecond range. This wide range of sensitivity makes EPR especially useful for studying protein dynamics. Depending on the rigidity of the attached spin label, it is possible to follow steric restrictions imposed by the environment of the label, motion of a segment of a polypeptide chain, and Brownian diffusion of a whole molecule [27]. We analyzed spectra according to the plot described by Mchaourab et al. [27], revealing characteristic patterns of spin-label mobility in relation to the secondary structures of proteins (including spin label on the surface of helices, interspherical loops, sites involved in tertiary contacts, and buried sites). Motional averaging of the spin label can be characterized by anisotropy averaging (the second moment of the spectrum) and by g-factor averaging (the central-field line width). We plotted the reciprocal of the splitting squared against the reciprocal of the central line width (see Materials and methods). Data from our
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Fig. 3. Secondary structure and tertiary contact predictions from a double reciprocal plot of the spin-label mobility of Tn complexes under various conditions. The reciprocal of the second moment of the spectrum ÆH2æ1 is plotted against the reciprocal of the central line width DH 1 0 . Spin-label mobility is used as an empirical method to evaluate secondary and tertiary structures of labeled sites including those on the helix surface and loop, in buried sites, and sites involved in tertiary contact. See Materials and methods for details. A and Tm indicate actin filaments and tropomyosin, respectively.
Fig. 2. EPR spectra of reconstituted filaments. Spectra of spin-labeled troponin complexes in reconstituted thin filaments in the +Ca2+ (A) or Ca2+ state (B), and those of spin-labeled TnI in reconstituted TnI–actin (C) and TnI–tropomyosin–actin filaments (D). In (B), the slow component with 28% of the total spins (thin curve) was produced by digital subtraction of the fast component (i.e., spectrum in (A) with 72% of the total spins (dashed curve) from observed spectra (solid curve) In (C), the fast component of TnI–actin (thin curve) was produced by subtracting 45% of the total spins of the slow component of TnC–I–T–tropomyosin– actin (Ca2+) (dashed curve) from the whole spectrum (solid curve). In (D), the slow component of TnI–tropomyosin–actin (thin curve) was produced by subtracting the fast component of TnI–actin containing 33% of the total spins (dashed curve) from the whole spectra (solid curve).
measurements were superimposed on the plot reported by Mchaourab et al. (Fig. 3) where data from T4 lysozyme were clustered and were characterized by four structurally different areas. Data from MSL–Tn complexes alone, and from reconstituted complexes on thin filaments in the +Ca2+ state, were both recovered on the same area clustering data from the spin label, at a helix surface. On the other hand, in the Ca2+ state, only data from troponin complexes were recovered on a loop-like area, whereas those from reconstituted filaments (TnI–actin, TnI–tropomyosin–actin, and TnC–I–T–tropomyosin–actin) were recovered on a tertiary contact area. We therefore postulated regarding the mechanism for the switch action of TnI. In troponin complexes in the
Ca2+ state, the C-terminal region of TnI is away from TnC, and flexible without interactions with other proteins, leading to the flexible structure of Cys133 that is like a loop or an independently tumbling helical segment. When Ca2+ is present, the Cys133 region of TnI folds into a helix or is stabilized by binding with the N-lobe of Ca2+-saturated TnC. These observations are consistent with the crystal structure of chicken skeletal troponin complexes as recently revealed by Vinogradova et al. [24]. Although the C-terminal region including Asn133 in TnI has been resolved to a loop in the +Ca2+ state, the switch segment (amino acids 113–131) very close to Asn133 forms an a-helix and interacts with the N-lobe of TnC. In the Ca2+ state, the switch segment and C-terminus including Asn133 in TnI are not resolved due to the low density. A recent NMR study [37] showed that 132–140 residues in the a-helix of TnI were flexible and mobile in solution. In reconstituted thin filaments, the second actin binding site near Cys133 binds to actin, and the attached spin label is involved in tertiary contact. Recently, Murakami et al. [37] showed that the C-terminal domain including Asn133 next to the switch segment bound to an outer domain of the actin subunit at low Ca2+ conditions. Most spectra from spin-labeled TnI on actin filaments have two components (Table 1). The moderately immobilized component (s = 4.8 ns) seen in the spectrum from reconstituted thin filaments in the Ca2+ state was almost identical to that in the spectrum observed in the +Ca2+ state and also from TnC–I–T complexes in the +Ca2+ state. The rigidly immobilized component (s = 45.2 ns) produced by spectral subtraction was recovered in the buried area according to the plot of Fig. 3 (data not shown).
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468
Therefore, it is possible that the Cys133 region is rigidly buried in the interface with actin in the actin-bound state, but it flutters in an equilibrium between TnC- and actinbound states on the thin filaments, even in the absence of Ca2+. The fast component (s = 2.6 ns) seen in spectra from TnI–actin or TnI–tropomyosin–actin was different from the motion observed in the thin filaments of TnC– I–T–tropomyosin–actin in the Ca2+ state (s = 4.8 ns), but was similar to that observed in TnC–I–T complexes in the Ca2+ state (s = 1.9 ns). Again, it is possible that the Cys133 region exists in equilibrium between an actinbound buried state and a freely dissociated state. These possibilities mean that TnC can bind a part of the switch segments of TnI even in the absence of Ca2+ (because it might be close enough to the actin sites for TnI to attain high affinity for the segments) while upon Ca2+ activation it binds all of the segments. We are now undertaking site-directed distance measurements using dipolar EPR methods [10,11] to identify the detailed mechanisms involved in interactions between the C-terminal region of TnI and the N-lobe of TnC in the +Ca2+ state, and with surfaces of tropomyosin and actin in the Ca2+ state.
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
Acknowledgments We thank Dr. M. Miki for valuable discussions and for guidance in respect to troponin preparation. This work was supported by grants from the Special Coordination Funds from the Ministry of Education, Culture, Sports, Science and Technology, Japan (to T.A.). References [1] S. Ebashi, M. Endo, Calcium ion and muscle contraction, Prog. Biophys. Mol. Biol. 18 (1968) 123–183. [2] S. Ebashi, M. Endo, I. Otsuki, Control of muscle contraction, Q. Rev. Biophys. 2 (1969) 351–384. [3] J. Gergely, Molecular switches in troponin, Adv. Exp. Med. Biol. 453 (1998) 169–176. [4] A.S. Zot, J.D. Potter, Structural aspects of troponin–tropomyosin regulation of skeletal–muscle contraction, Annu. Rev. Biophys. Biophys. Chem. 16 (1987) 535–559. [5] M.X. Li, X. Wang, B.D. Sykes, Structural based insights into the role of troponin in cardiac muscle pathophysiology, J. Muscle Res. Cell Motil. 25 (2004) 559–579. [6] R.L. Moss, M. Razumova, D.P. Fitzsimons, Myosin crossbridge activation of cardiac thin filaments: implications for myocardial function in health and disease, Circ. Res. 94 (2004) 1290–1300. [7] O. Herzberg, J. Moult, M.N.G. James, A model for the Ca2+induced conformational transition of troponin C, J. Biol. Chem. 261 (1986) 2638–2679. [8] C.M. Slupsky, B.D. Sykes, NMR solution structure of calciumsaturated skeletal muscle troponin C, Biochemistry 34 (1995) 15953– 15964. [9] W.J. Dong, J. Xing, M. Villain, M. Hellinger, J.M. Robinson, M. Chandra, R.J. Solaro, P.K. Umeda, H.C. Cheung, Conformation of the regulatory domain of cardiac muscle troponin C in its complex with cardiac troponin I, J. Biol. Chem. 274 (1999) 31382–31390. [10] S. Ueki, M. Nakamura, T. Komori, T. Arata, Site-directed spin labeling electron paramagnetic resonance study of the calcium-induced struc-
[19]
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
467
tural transition in the N-domain of human cardiac troponin C complexed with troponin I, Biochemistry 44 (2005) 411–416. M. Nakamura, S. Ueki, H. Hara, T. Arata, Calcium structural transition of human cardiac troponin C in reconstituted muscle fibres as studied by site-directed spin labelling, J. Mol. Biol. 348 (2005) 127– 137. B.A. Levine, A.J. Moir, S.V. Perry, The interaction of troponin-I with the N-terminal region of actin, Eur. J. Biochem. 172 (1988) 389–397. J.A. Talbot, R.S. Hodges, Comparative studies on the inhibitory region of selected species of troponin-I. The use of synthetic peptide analogs to probe structure–function relationships, J. Biol. Chem. 256 (1981) 12374–12378. C.S. Farah, C.A. Miyamoto, C.H. Ramos, A.C. da Silva, R.B. Quaggio, K. Fujimori, L.B. Smillie, F.C. Reinach, Structural and regulatory functions of the NH2- and COOH-terminal regions of skeletal muscle troponin I, J. Biol. Chem. 269 (1994) 5230–5240. H.M. Rarick, X.H. Tu, R.J.A.F. Martin, The C terminus of cardiac troponin I is essential for full inhibitory activity and Ca2+ sensitivity of rat myofibrils, J. Biol. Chem. 272 (1997) 26887–26892. B. Tripet, J.E. Van Eyk, R.S. Hodges, Mapping of a second actin– tropomyosin and a second troponin C binding site within the C terminus of troponin I, and their importance in the Ca2+-dependent regulation of muscle contraction, J. Mol. Biol. 271 (1997) 728–750. C.H. Ramos, Mapping subdomains in the C-terminal region of troponin I involved in its binding to troponin C and to thin filament, J. Biol. Chem. 274 (1999) 18189–18195. Y. Luo, J. Leszyk, Y. Qian, J. Gergely, T. Tao, Residues 48 and 82 at the N-terminal hydrophobic pocket of rabbit skeletal muscle troponin-C photo-cross-link to Met121 of troponin-I, Biochemistry 38 (1999) 6678–6688. T. Tao, E. Gowell, G.M. Strasburg, J. Gergely, P.C. Leavis, Ca2+ dependence of the distance between Cys-98 of troponin C and Cys133 of troponin I in the ternary troponin complex. Resonance energy transfer measurements, Biochemistry 28 (1989) 5902–5908. T. Tao, B.J. Gong, P.C. Leavis, Calcium-induced movement of troponin-I relative to actin in skeletal muscle thin filaments, Science 247 (1990) 1339–1341. M. Miki, Resonance energy transfer between points in a reconstituted skeletal muscle thin filament. A conformational change of the thin filament in response to a change in Ca2+ concentration, Eur. J. Biochem. 187 (1990) 155–162. Y. Shitaka, C. Kimura, T. Iio, M. Miki, Kinetics of the structural transition of muscle thin filaments observed by fluorescence resonance energy transfer, Biochemistry 43 (2004) 10739–10747. S. Takeda, A. Yamashita, K. Maeda, Y. Maeda, Structure of the core domain of human cardiac troponin in the Ca(2+)-saturated form, Nature 424 (2003) 35–41. M.V. Vinogradova, D.B. Stone, G.G. Malanina, C. Karatzaferi, R. Cooke, R.A. Mendelson, R.J. Fletterick, Ca(2+)-regulated structural changes in troponin, Proc. Natl. Acad. Sci. USA 102 (2005) 5038–5043. W.L. Hubbell, A. Gross, R. Langen, M.A. Lietzow, Recent advances in site-directed spin labeling of proteins, Curr. Opin. Struct. Biol. 8 (1998) 649–656. W.L. Hubbell, D.S. Cafiso, C. Altenbach, Identifying conformational changes with site-directed spin labeling, Nat. Struct. Biol. 7 (2000) 735–739. H.S. Mchaourab, M.A. Lietzow, K. Hideg, W.L. Hubbell, Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics, Biochemistry 35 (1996) 7692–7704. S. Onishi, K. Maruyama, S. Ebashi, Calcium-induced conformational changes and mutual interactions of troponin components as studied by spin labeling, J. Biochem. 78 (1975) 73–81. J.A. Spudich, S. Watt, The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin–troponin complex with actin and the proteolytic fragments of myosin, J. Biol. Chem. 246 (1971) 4866–4871.
468
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468
[30] S. Ebashi, A. Kodama, F. Ebashi, Troponin. I. Preparation and physiological function, J. Biochem. (Tokyo) 64 (1968) 465– 477. [31] P.C.S. Chong, R.S. Hodges, Proximity of sulfhydryl groups to the sites of interaction between components of the troponin complex from rabbit skeletal muscle, J. Biol. Chem. 257 (1982) 2549–2555. [32] T. Ojima, K. Nishita, Separation of Akazara scallop and rabbit troponin components by a single-step chromatography on CMToyopearl, J. Biochem. (Tokyo) 104 (1988) 9–11. [33] N. Matsuo, Y. Nagata, J. Nakamura, T. Yamamoto, Coupling of calcium transport with ATP hydrolysis in scallop sarcoplasmic reticulum, J. Biochem. (Tokyo) 131 (2002) 375–381.
[34] A. Redfield, The theory of relaxation processes, Adv. Magn. Reson. 1 (1965) 1. [35] D.E. Budil, S. Lee, S. Saxena, J.H. Freed, Non-linear least squares analysis of slow-motion EPR spectra in one and two dimensions using a modified Levenberg–Marquardt algorithm, J. Magn. Reson. 120 (1996) 155–189. [36] H.C. Li, K. Hideg, P.G. Fajer, The mobility of troponin C and troponin I in muscle, J. Mol. Recognit. 10 (1997) 194–201. [37] K. Murakami, F. Yumoto, S.Y. Ohki, T. Yasunaga, M. Tanokura, T. Wakabayashi, Structural basis for Ca2+-regulated muscle relaxation at interaction sites of troponin with actin and tropomyosin, J. Mol. Biol. 352 (2005) 178–201.
Calcium-dependent movement of troponin I between troponin C and actin as revealed by spin-labeling EPR Tomoki Aihara, Shoji Ueki, Motoyoshi Nakamura, Toshiaki Arata
*
Department of Biological Sciences, Graduate School of Science, Osaka University and CREST/JST, Toyonaka, Osaka 560-0043, Japan Received 17 November 2005 Available online 15 December 2005
Abstract We measured EPR spectra from a spin label on the Cys133 residue of troponin I (TnI) to identify Ca2+-induced structural states, based on sensitivity of spin-label mobility to flexibility and tertiary contact of a polypeptide. Spectrum from Tn complexes in the Ca2+ state showed that Cys133 was located at a flexible polypeptide segment (rotational correlation time s = 1.9 ns) that was free from TnC. Spectra of both Tn complexes alone and those reconstituted into the thin filaments in the +Ca2+ state showed that Cys133 existed on a stable segment (s = 4.8 ns) held by TnC. Spectra of reconstituted thin filaments (Ca2+ state) revealed that slow mobility (s = 45 ns) was due to tertiary contact of Cys133 with actin, because the same slow mobility was found for TnI–actin and TnI–tropomyosin–actin filaments lacking TnC, T or tropomyosin. We propose that the Cys133 region dissociates from TnC and attaches to the actin surface on the thin filaments, causing muscle relaxation at low Ca2+ concentrations. 2005 Elsevier Inc. All rights reserved. Keywords: Muscle regulation; Actin; Troponin; Tropomyosin; Spin labeling; EPR
Contraction of vertebrate striated muscles is regulated by troponin (Tn)/tropomyosin complexes located on the actin thin filaments [1–6]. Troponin is a calcium binding protein, and its sensitivity to calcium concentration is important for its regulation. Tn is composed of three heterotrimeric subunits, i.e., a calcium (Ca2+) binding subunit (TnC), an inhibitory subunit (TnI), and a tropomyosinbinding subunit (TnT). The crystal structure of TnC suggested that the mechanism underlying its activation involved Ca2+ binding to two regulatory sites, inducing movement of helices in the N-terminal domain, and so exposing a hydrophobic patch and further augmenting interaction with TnI [7]. This mechanism was consistent with that described in other structural studies [8–11]. TnI contains a few functional regions at the C-terminus. Its inhibitory segment (amino acids 104–115) and second actin-binding site (amino acids 140–148) can bind to actin and inhibit actomyosin ATPase activity [12–17]. Moreover, *
Corresponding author. Fax: +81 6 6850 5441. E-mail address: [email protected] (T. Arata).
0006-291X/$ - see front matter 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2005.12.030
the switch segment (amino acids 116–131) between two actin-binding sites is thought to interact with the N-lobe of TnC in the presence of Ca2+ [18] and to dissociate from it by binding the second site to actin in the absence of Ca2+. Using fluorescence resonance energy transfer (FRET), a number of studies showed that Cys133 near the TnI switch segment underwent Ca2+-dependent movement around TnC and actin subunits on the thin filaments [19–22]. Crystal structures of cardiac and skeletal TnC/TnI/TnT complexes have been solved, and showed that the switch segment of TnI was clutched by the N-lobe of TnC in the +Ca2+ state [23,24], but was disordered in the Ca2+ state [24]. Therefore, it is important to understand how Tn complexes undergo Ca2+-triggered structural changes on the thin filaments and in muscle fibers. Site-directed spin-labeling electron paramagnetic resonance (SDSL-EPR) spectroscopy has been used to determine secondary, tertiary, and quaternary structures, and associated conformational changes of proteins [25,26]. Depending on the rigidity of the attached spin label, it is possible to investigate steric restrictions imposed by the environment
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of the label, motion of a segment of a polypeptide chain, or Brownian diffusion of a whole molecule [27]. Onishi et al. [28] reported that a spin label, presumably positioned on Cys133 in TnI, showed Ca2+ -dependent immobilization in Tn complexes. Here, we confirmed that removal of Ca2+ increased the mobility of the side-chain spin label on Cys133, located near the switch segment of the rabbit TnI complexes in solution. According to the mobility–structure relationships in standard proteins [27], detection of a mobile spectrum arises from a loop-like flexible structure, while that of an immobile spectrum arises from a spin label on a stable loop or a helix surface. With the reconstituted thin filaments, Ca2+ removal resulted in opposite effects, i.e., it induced rigid immobilization of the spin labels, whereas the spectrum in the presence of Ca2+ was the same as that in Tn complexes. The rigidly immobilized component of spin labels was also observed in reconstituted TnI–actin and TnI–tropomyosin–actin filaments. This suggests that the side chain of Cys133 makes tertiary contact with the actin surface after forming a flexible structure free from TnC, upon Ca2+ removal. Therefore, we propose that the switch segment undergoes a Ca2+-dependent movement between TnC and actin on the thin filaments. Materials and methods Reagents. 4-Maleimido-2,2,6,6-tetramethyl-1-piperidinoxy (MSL) was from Sigma Chemicals. The BCA–protein assay reagents used to measure protein concentrations were from Pierce Chemicals. Other reagents were of analytical grade. Protein preparation. Rabbit muscle protein was prepared from back and leg muscles, and chicken from breast muscles. Actin was prepared from acetone powder of rabbit skeletal muscles, according to the method of Spudich and Watt [29]. Native troponin complexes (TnC–I–T complexes) and tropomyosin were prepared from rabbit and chicken skeletal muscle residues after extraction of myosin by the method of Ebashi et al. [30]. After dialysis with 20 mM Tris–HCl (pH 7.5), 30 mM KCl, 0.1 mM CaCl2 (buffer A), and 1 mM DTT, troponin was purified using a Q-Sepharose Fast Flow column (2.5 · 10 cm) and eluted with a linear 30–400 mM KCl gradient. Fractions containing TnC–I–T complexes were identified, collected by measuring absorption at 280 nm and by SDS– PAGE, and finally dialyzed with buffer A containing 1 mM DTT. Spin labeling of troponin complexes. Prior to spin labeling, purified troponin complexes were passed through a Sephadex G-25 column (1 · 20 cm) equilibrated with buffer A without DTT. Then, troponin (10– 40 lM) was reacted with an equimolar amount of MSL overnight in the dark at 4 C. The following day, unreacted MSL was removed by passing labeled troponin (MSL-Tn) through a Sephadex G-25 column equilibrated with buffer A. When removing DTT or unreacted MSL for large-scale preparations (>20 ml), dialysis using buffer A was done instead of using a Sephadex G-25 column. For EPR measurements, the MSL-labeled troponin solution was concentrated up to 1–3 mg/ml (10–40 lM) and examined after 0.1 mM CaCl2 or 1 mM EGTA was added in buffer A to achieve the +Ca2+ state or Ca2+ state, respectively. MSL-labeled TnI (MSL–TnI) was isolated from MSL–Tn complexes by the method of Ojima and Nishita [32]. MSL–Tn was dialyzed against 6 M urea, 30 mM KCl, 1 mM EDTA, and 10 mM Tris–HCl (pH 7.6), and applied to a CM-Toyopearl 650 M column (2.5 · 10 cm). Following preequilibration with the same buffer (to wash out isolated TnC), MSL–TnI was eluted with a linear gradient of 30–300 mM KCl, before eluting TnT. Fractions containing MSL–TnI were identified by measuring absorption at 280 nm and by SDS–PAGE. Eluted MSL–TnI was gently dialyzed against 100 mM KCl and 20 mM Tris–HCl (pH 7.5). After insoluble
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components were removed by centrifugation, the MSL–TnI preparation was used for reconstitution of thin (TnC–I–T-tropomyosin–actin) Tn-actin and TnI–tropomyosin–actin filaments. Reconstitution of thin filaments G-actin was incubated in 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, and 4 mM MgCl2 for 30 min. Tropomyosin and MSL–Tn were added to G-actin solution (ca. 100 lM, 0.3–0.5 ml), in the presence of 1 mM CaCl2 (+Ca2+ state) or 5 mM EGTA (Ca2+ state), and further incubated for 1 h. Molar ratio was set as 1:1:3 for tropomyosin, MSL–Tn complexes, and actin. Then, the mixture was centrifuged at 100,000g for 30 min, and the pellet co-sedimented with MSL–Tn was finally resuspended in a small volume (20–100 ll) of 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, 4 mM MgCl2, and 1 mM CaCl2 or 5 mM EGTA. Amounts of TnC, TnI, TnT, tropomyosin, and actin were assessed by SDS–PAGE. The suspension, containing 20–40 lM MSL–troponin, was used for EPR measurements. TnI–actin and TnI–tropomyosin–actin were also prepared as described above. After G-actin (ca. 100 lM, 0.3–0.5 ml) was incubated in 100 mM KCl and 4 mM MgCl2 for 30 min, tropomyosin was mixed with actin solution at a molar ratio of 1:3 with G-actin. Isolated MSL–TnI was mixed with actin or actin–tropomyosin solution at a molar ratio of 1:3 with G-actin and incubated for 1 h. The mixture was centrifuged at 100,000g for 30 min, and the pellet co-sedimented with MSL–TnI was finally resuspended in a small volume (ca. 20 ll) of 20 mM imidazole–HCl (pH 7.0), 100 mM KCl, and 4 mM MgCl2. Amounts of TnI, tropomyosin, and actin were assessed by SDS–PAGE. The suspension, containing 20– 40 lM MSL–TnI, was used for EPR measurements. ATPase assay. Inhibitory activity of troponin was measured using an actomyosin ATPase assay. Troponin (1–4 lM) was mixed with 1 lM tropomyosin, 3.5 lM actin, and 0.6 lM myosin in 0.1 M KCl, 4 mM MgCl2, 1 mM CaCl2, and 20 mM Tris–HCl (pH 7.5). The ATPase reaction was coupled with a regeneration system as described by Matsuo et al. [33], where 2 mM phosphoenolpyruvic acid, 0.3 mM NADH, 38 U/ ml pyruvate kinase, and 50 U/ml lactate dehydrogenase were mixed. The reaction was started at 25 C by addition of 100 lM ATP, and time course of NADH absorption at 340 nm was followed for 10 min. Next, 5–10 mM EGTA was mixed with the sample and followed for 10 min. Velocities of the decrease in absorption were compared on charts according to the methods of Nakamura et al. [11] and were defined as: [ATPase (+Ca2+) activity ATPase(Ca2+) activity]/ATPase (+Ca2+) activity. EPR measurements and analysis EPR spectroscopy was performed using a Bruker ELEXSYS E500 spectrometer equipped with a dielectric resonator. Sample solutions (ca. 15 ll) were loaded into capillaries (inside diameter, 1.0 mm), inserted into the resonator, and EPR spectra acquired using 1-G field modulation amplitude at 100 kHz and 5-mW incident microwave power at 25 C. The rotational correlation time (s) was determined for the original spectrum and for those of fast or slow components produced by computer subtraction. The rotational correlation time in the fast motion regime (s 109 s) was calculated according to the equation of Redfield [34]: s = 1.19 · 109 DH0{(V0/V+1)0.5 + (V0/V1)0.52}, where DH0 is the peak-to-peak line width of the central line and Vm is the peak-to-peak height [34]. The rotational correlation time in the slow motion regime (s 109 2 · 107 s) was calculated according to an equation from Freed and co-workers [35]: s = a (1Teff/Tmax)b, where a = 5.4 · 1010 s, b = 1.36, Tmax (=35 G) is the rigid limit for a particular spin, and 2Teff is the width between the low-field and high-field absorption peaks [35]. Further, we plotted the reciprocal second moment ÆHæ2 (G2) versus the reciprocal central line width ÆDH0æ calculated from EPR spectra, according to Mchaourab et al. [27]. The following equation was used: P P hHi2 ¼ Ai DB2i = Ai , where Ai is the intensity at i field on the absorption spectrum and DBi (G) is the absolute value at i field assessed
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from the peak of the central extreme. We also characterized the secondary structure where a spin label existed.
Results and discussion Spin labeling of Cys133 on TnI We used SDSL-EPR spectroscopy to determine conformational changes in the C-terminal region of TnI in troponin complexes. Rabbit skeletal TnI was exclusively labeled with MSL because Cys133 near the switch segment of TnI was the most reactive cysteine in the complex state as reported by Chong and Hodges [31]. We investigated the effects of spin labeling. Actomyosin ATPase of MSL-Tn inhibited 85–95% of troponin labeling, indicating that spin labeling did not significantly affect inhibition by actomyosin ATPase. Labeling efficiency for rabbit troponin, estimated from double integration of the spectrum, was more than 0.8 mol/mol troponin complexes. For chicken troponin where Cys133 was substituted with another amino acid residue, efficiency was about 0.5–0.6 mol/mol troponin complexes. The lower reactivity for the other cysteines of chicken troponin suggested that for rabbits the fraction of spin-labeled cysteines other than Cys133 was very small [31,36]. Mobility of the spin label attached to Cys133 of TnI in troponin C–I–T complexes We measured EPR spectra from the spin label (MSL) of Cys133 of TnI in ternary TnC–I–T complexes to investigate conformational changes of the switch segment in
TnI, induced by Ca2+. Fig. 1 (left) shows spectra of rabbit skeletal troponin complexes in the presence and absence of Ca2+. In the +Ca2+ state (top), absorption line shapes were broad, and the two extremes at the low- and high-field were small relative to the central peak. From the spectral parameters DH0 = 3.0–3.4, V0/V1 = 7.0–13.0, and V0/V+1 = 2.1–2.4 (see Materials and methods), we deduced a slow motion of rotational correlation time s = 4.80 ± 1.57 ns (Table 1). In the Ca2+ state (1 mM EGTA), sharp peaks (at ca. 3420 G) appeared with a very small broad peak (at ca. 3410 G). Similar changes were seen at high magnetic field (at ca. 3450 G). Sharp major peaks (DH0 = 2.4–2.6, V0/V1 = 3.9–6.2, and V0/V+1 = 1.3–1.6) corresponded to a correlation time of s = 1.93 ± 0.57 ns (Table 1). These spectral changes were almost the same as those reported by Onishi et al. [28]. Excess Ca2+ (final concentration, 10 mM) was added to achieve the +Ca2+ state, and the spectrum returned to the original one, indicating a reversible reaction. Similar results were obtained for binary complexes of MSL-labeled TnI and TnC (Table 1). Chicken skeletal troponin was also prepared and measured under the same conditions. In chicken TnI, asparagine was substituted for Cys133 near the switch segment. Fig. 1 shows spectra of rabbit troponin (left) compared with those of chicken (right). Chicken spectra showed weak signal intensity compared with those of rabbit. Broad peaks (2Teff = ca. 63.0 G) at ca. 3410 and ca. 3470 G indicated rigid immobilization (s = 12–13 ns). Similar but very small peaks (at ca. 3410 and ca. 3470 G) in the slow motion regime were seen from rabbit troponin in the Ca2+ state but not in the +Ca2+ state. Spectra were not sensitive to Ca2+ concentration. Therefore, most of the spin labels were
Fig. 1. EPR spectra of spin-labeled troponin complexes from rabbit (left) and chicken (right) skeletal muscles in the +Ca2+ (top) and Ca2+ (bottom) states.
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468 Table 1 Summary of the rotational correlation time (s) of the spin label in TnI of rabbit s (ns)
TnC–I TnC–I–T TnC–I–T–tropomyosin–actin TnI–actin TnI–tropomyosin–actin
Ca2+ +Ca2+ Ca2+ +Ca2+ Ca2+ +Ca2+
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Cys64 of TnI; and Cys98 of TnC) did not result in mobility changes of spin labels attached on the thin filaments. MSL–TnI reconstituted with actin and tropomyosin–actin
Fast
Slow
1.67 3.26 1.93 ± 0.57a 4.80 ± 1.57a 4.76c (72%)b 4.76 2.61c (55%) 2.61c (33%)
— — — — 45.2c (28%) — 45.2c (45%) 43.0c (67%)
a
Values are means ± standard deviation for three preparations. Values in parentheses are the populations of the fast and slow components. c Rotational correlation times were determined for the spectra produced by spectral subtraction. The slow component of TnC–I–T–tropomyosin– actin (Ca2+) was obtained by digital subtraction of the spectrum of TnC–I–T–tropomyosin–actin (+Ca2+) containing 72% of the total spins from the whole spectrum. The fast component of TnI–actin was produced by subtracting 45% of the total spins of the slow component of TnC–I–T– tropomyosin–actin (Ca2+) from the whole spectrum. The slow component of TnI–tropomyosin–actin was produced by subtracting the fast component of TnI–actin containing 33% of the total spins from the whole spectrum. b
used to selectively label Cys133 of rabbit TnI and showed mobility changes induced by Ca2+ while spin labels of any other cysteines (presumably Cys48 and Cys64 of TnI; and Cys 98 of TnC) did not result in any spectral changes. MSL–TnI in reconstituted thin filaments To examine how spin-label mobility of Cys133 in TnI changes in response to Ca2+ on thin filaments, we reconstituted them with MSL–Tn–tropomyosin, and actin. Fig. 2 shows EPR spectra from rabbit skeletal thin filaments in the presence (A) and absence (B) of Ca2+. In the +Ca2+ state, spectrum shape mimicked that of troponin C–I–T complexes alone. The rotational correlation time was 4.72 ns (Table 1). On the other hand, in the Ca2+ state, a marked shoulder appeared at the lower field (about 3410 G). Similar corresponding changes were seen at a higher magnetic field (3450 G). The shoulder indicates a component of slow motion. The rotational correlation time was calculated as 45.2 ns from 2Teff (67.3 G) of the spectrum produced by spectral subtraction of the spectrum of +Ca2+ state containing 72% of the total spins from that of the Ca2+ state. We could not observe any enhancement of the fast motional component by adding EGTA, unlike MSL–Tn alone. These results were very similar to those observed for MSL–TnI reconstituted into myofibrils [37]. The experiments with the thin filaments were also performed using chicken troponin complexes. Spin-labeled chicken TnC–I–T complexes and rabbit actin were mixed and reconstituted into thin filaments. The resulting spectrum was the same as that of TnC–I–T complexes alone and revealed no Ca2+ effects (data not shown). This indicated that cysteines other than Cys133 (presumably Cys48 and
To examine whether Ca2+-removal induced immobilization of the spin label on Cys133 of TnI in thin filaments, and that it was due to the binding of the side chain of Cys133 to actin or TnT/C, we reconstituted isolated MSL–TnI with actin with or without tropomyosin. Figs. 2C and D show spectra from TnI–actin and TnI–tropomyosin–actin, respectively. No Ca2+ effect was observed, unlike in the thin filaments (TnC–I–T–tropomyosin–actin). Both spectra, especially from TnI–tropomyosin–actin, were very similar to that from thin filaments reconstituted in the Ca2+ state. Both spectra consisted of two distinct components, i.e., fast and slow motions. The slow component (2Teff = 67.3 G) was close to the motion observed in thin filaments in the Ca2+ state (s = 45 ns). Therefore, the fast component of TnI–actin was easily produced by subtracting 45% of the total spins of the slow component obtained by decomposition of the two spectra of TnC–I– T–tropomyosin–actin shown in Figs. 2A and B. From the spectral parameters DH0 = 3.1, V0/V1 = 4.5, and V0/ V+1 = 1.7, we deduced rotational correlation time s = 2.6 ns for the fast component (Table 1). This fast component containing 33% of the total spins was subtracted from the spectrum of TnI–tropomyosin–actin (Fig. 2D) to produce the same slow component again (2Teff = 67.2 G, s = 43 ns). These results were a strong indication that Cys133 in TnI interacted directly with actin in thin filaments in the Ca2+ state, because it could interact with actin in the same way, irrespective of troponin or tropomyosin. Secondary structure prediction from side-chain mobility of the spin label Electron paramagnetic resonance (EPR) spectroscopy allows the investigation of motional sensitivity within a nanosecond to submillisecond range. This wide range of sensitivity makes EPR especially useful for studying protein dynamics. Depending on the rigidity of the attached spin label, it is possible to follow steric restrictions imposed by the environment of the label, motion of a segment of a polypeptide chain, and Brownian diffusion of a whole molecule [27]. We analyzed spectra according to the plot described by Mchaourab et al. [27], revealing characteristic patterns of spin-label mobility in relation to the secondary structures of proteins (including spin label on the surface of helices, interspherical loops, sites involved in tertiary contacts, and buried sites). Motional averaging of the spin label can be characterized by anisotropy averaging (the second moment of the spectrum) and by g-factor averaging (the central-field line width). We plotted the reciprocal of the splitting squared against the reciprocal of the central line width (see Materials and methods). Data from our
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Fig. 3. Secondary structure and tertiary contact predictions from a double reciprocal plot of the spin-label mobility of Tn complexes under various conditions. The reciprocal of the second moment of the spectrum ÆH2æ1 is plotted against the reciprocal of the central line width DH 1 0 . Spin-label mobility is used as an empirical method to evaluate secondary and tertiary structures of labeled sites including those on the helix surface and loop, in buried sites, and sites involved in tertiary contact. See Materials and methods for details. A and Tm indicate actin filaments and tropomyosin, respectively.
Fig. 2. EPR spectra of reconstituted filaments. Spectra of spin-labeled troponin complexes in reconstituted thin filaments in the +Ca2+ (A) or Ca2+ state (B), and those of spin-labeled TnI in reconstituted TnI–actin (C) and TnI–tropomyosin–actin filaments (D). In (B), the slow component with 28% of the total spins (thin curve) was produced by digital subtraction of the fast component (i.e., spectrum in (A) with 72% of the total spins (dashed curve) from observed spectra (solid curve) In (C), the fast component of TnI–actin (thin curve) was produced by subtracting 45% of the total spins of the slow component of TnC–I–T–tropomyosin– actin (Ca2+) (dashed curve) from the whole spectrum (solid curve). In (D), the slow component of TnI–tropomyosin–actin (thin curve) was produced by subtracting the fast component of TnI–actin containing 33% of the total spins (dashed curve) from the whole spectra (solid curve).
measurements were superimposed on the plot reported by Mchaourab et al. (Fig. 3) where data from T4 lysozyme were clustered and were characterized by four structurally different areas. Data from MSL–Tn complexes alone, and from reconstituted complexes on thin filaments in the +Ca2+ state, were both recovered on the same area clustering data from the spin label, at a helix surface. On the other hand, in the Ca2+ state, only data from troponin complexes were recovered on a loop-like area, whereas those from reconstituted filaments (TnI–actin, TnI–tropomyosin–actin, and TnC–I–T–tropomyosin–actin) were recovered on a tertiary contact area. We therefore postulated regarding the mechanism for the switch action of TnI. In troponin complexes in the
Ca2+ state, the C-terminal region of TnI is away from TnC, and flexible without interactions with other proteins, leading to the flexible structure of Cys133 that is like a loop or an independently tumbling helical segment. When Ca2+ is present, the Cys133 region of TnI folds into a helix or is stabilized by binding with the N-lobe of Ca2+-saturated TnC. These observations are consistent with the crystal structure of chicken skeletal troponin complexes as recently revealed by Vinogradova et al. [24]. Although the C-terminal region including Asn133 in TnI has been resolved to a loop in the +Ca2+ state, the switch segment (amino acids 113–131) very close to Asn133 forms an a-helix and interacts with the N-lobe of TnC. In the Ca2+ state, the switch segment and C-terminus including Asn133 in TnI are not resolved due to the low density. A recent NMR study [37] showed that 132–140 residues in the a-helix of TnI were flexible and mobile in solution. In reconstituted thin filaments, the second actin binding site near Cys133 binds to actin, and the attached spin label is involved in tertiary contact. Recently, Murakami et al. [37] showed that the C-terminal domain including Asn133 next to the switch segment bound to an outer domain of the actin subunit at low Ca2+ conditions. Most spectra from spin-labeled TnI on actin filaments have two components (Table 1). The moderately immobilized component (s = 4.8 ns) seen in the spectrum from reconstituted thin filaments in the Ca2+ state was almost identical to that in the spectrum observed in the +Ca2+ state and also from TnC–I–T complexes in the +Ca2+ state. The rigidly immobilized component (s = 45.2 ns) produced by spectral subtraction was recovered in the buried area according to the plot of Fig. 3 (data not shown).
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468
Therefore, it is possible that the Cys133 region is rigidly buried in the interface with actin in the actin-bound state, but it flutters in an equilibrium between TnC- and actinbound states on the thin filaments, even in the absence of Ca2+. The fast component (s = 2.6 ns) seen in spectra from TnI–actin or TnI–tropomyosin–actin was different from the motion observed in the thin filaments of TnC– I–T–tropomyosin–actin in the Ca2+ state (s = 4.8 ns), but was similar to that observed in TnC–I–T complexes in the Ca2+ state (s = 1.9 ns). Again, it is possible that the Cys133 region exists in equilibrium between an actinbound buried state and a freely dissociated state. These possibilities mean that TnC can bind a part of the switch segments of TnI even in the absence of Ca2+ (because it might be close enough to the actin sites for TnI to attain high affinity for the segments) while upon Ca2+ activation it binds all of the segments. We are now undertaking site-directed distance measurements using dipolar EPR methods [10,11] to identify the detailed mechanisms involved in interactions between the C-terminal region of TnI and the N-lobe of TnC in the +Ca2+ state, and with surfaces of tropomyosin and actin in the Ca2+ state.
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
Acknowledgments We thank Dr. M. Miki for valuable discussions and for guidance in respect to troponin preparation. This work was supported by grants from the Special Coordination Funds from the Ministry of Education, Culture, Sports, Science and Technology, Japan (to T.A.). References [1] S. Ebashi, M. Endo, Calcium ion and muscle contraction, Prog. Biophys. Mol. Biol. 18 (1968) 123–183. [2] S. Ebashi, M. Endo, I. Otsuki, Control of muscle contraction, Q. Rev. Biophys. 2 (1969) 351–384. [3] J. Gergely, Molecular switches in troponin, Adv. Exp. Med. Biol. 453 (1998) 169–176. [4] A.S. Zot, J.D. Potter, Structural aspects of troponin–tropomyosin regulation of skeletal–muscle contraction, Annu. Rev. Biophys. Biophys. Chem. 16 (1987) 535–559. [5] M.X. Li, X. Wang, B.D. Sykes, Structural based insights into the role of troponin in cardiac muscle pathophysiology, J. Muscle Res. Cell Motil. 25 (2004) 559–579. [6] R.L. Moss, M. Razumova, D.P. Fitzsimons, Myosin crossbridge activation of cardiac thin filaments: implications for myocardial function in health and disease, Circ. Res. 94 (2004) 1290–1300. [7] O. Herzberg, J. Moult, M.N.G. James, A model for the Ca2+induced conformational transition of troponin C, J. Biol. Chem. 261 (1986) 2638–2679. [8] C.M. Slupsky, B.D. Sykes, NMR solution structure of calciumsaturated skeletal muscle troponin C, Biochemistry 34 (1995) 15953– 15964. [9] W.J. Dong, J. Xing, M. Villain, M. Hellinger, J.M. Robinson, M. Chandra, R.J. Solaro, P.K. Umeda, H.C. Cheung, Conformation of the regulatory domain of cardiac muscle troponin C in its complex with cardiac troponin I, J. Biol. Chem. 274 (1999) 31382–31390. [10] S. Ueki, M. Nakamura, T. Komori, T. Arata, Site-directed spin labeling electron paramagnetic resonance study of the calcium-induced struc-
[19]
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
467
tural transition in the N-domain of human cardiac troponin C complexed with troponin I, Biochemistry 44 (2005) 411–416. M. Nakamura, S. Ueki, H. Hara, T. Arata, Calcium structural transition of human cardiac troponin C in reconstituted muscle fibres as studied by site-directed spin labelling, J. Mol. Biol. 348 (2005) 127– 137. B.A. Levine, A.J. Moir, S.V. Perry, The interaction of troponin-I with the N-terminal region of actin, Eur. J. Biochem. 172 (1988) 389–397. J.A. Talbot, R.S. Hodges, Comparative studies on the inhibitory region of selected species of troponin-I. The use of synthetic peptide analogs to probe structure–function relationships, J. Biol. Chem. 256 (1981) 12374–12378. C.S. Farah, C.A. Miyamoto, C.H. Ramos, A.C. da Silva, R.B. Quaggio, K. Fujimori, L.B. Smillie, F.C. Reinach, Structural and regulatory functions of the NH2- and COOH-terminal regions of skeletal muscle troponin I, J. Biol. Chem. 269 (1994) 5230–5240. H.M. Rarick, X.H. Tu, R.J.A.F. Martin, The C terminus of cardiac troponin I is essential for full inhibitory activity and Ca2+ sensitivity of rat myofibrils, J. Biol. Chem. 272 (1997) 26887–26892. B. Tripet, J.E. Van Eyk, R.S. Hodges, Mapping of a second actin– tropomyosin and a second troponin C binding site within the C terminus of troponin I, and their importance in the Ca2+-dependent regulation of muscle contraction, J. Mol. Biol. 271 (1997) 728–750. C.H. Ramos, Mapping subdomains in the C-terminal region of troponin I involved in its binding to troponin C and to thin filament, J. Biol. Chem. 274 (1999) 18189–18195. Y. Luo, J. Leszyk, Y. Qian, J. Gergely, T. Tao, Residues 48 and 82 at the N-terminal hydrophobic pocket of rabbit skeletal muscle troponin-C photo-cross-link to Met121 of troponin-I, Biochemistry 38 (1999) 6678–6688. T. Tao, E. Gowell, G.M. Strasburg, J. Gergely, P.C. Leavis, Ca2+ dependence of the distance between Cys-98 of troponin C and Cys133 of troponin I in the ternary troponin complex. Resonance energy transfer measurements, Biochemistry 28 (1989) 5902–5908. T. Tao, B.J. Gong, P.C. Leavis, Calcium-induced movement of troponin-I relative to actin in skeletal muscle thin filaments, Science 247 (1990) 1339–1341. M. Miki, Resonance energy transfer between points in a reconstituted skeletal muscle thin filament. A conformational change of the thin filament in response to a change in Ca2+ concentration, Eur. J. Biochem. 187 (1990) 155–162. Y. Shitaka, C. Kimura, T. Iio, M. Miki, Kinetics of the structural transition of muscle thin filaments observed by fluorescence resonance energy transfer, Biochemistry 43 (2004) 10739–10747. S. Takeda, A. Yamashita, K. Maeda, Y. Maeda, Structure of the core domain of human cardiac troponin in the Ca(2+)-saturated form, Nature 424 (2003) 35–41. M.V. Vinogradova, D.B. Stone, G.G. Malanina, C. Karatzaferi, R. Cooke, R.A. Mendelson, R.J. Fletterick, Ca(2+)-regulated structural changes in troponin, Proc. Natl. Acad. Sci. USA 102 (2005) 5038–5043. W.L. Hubbell, A. Gross, R. Langen, M.A. Lietzow, Recent advances in site-directed spin labeling of proteins, Curr. Opin. Struct. Biol. 8 (1998) 649–656. W.L. Hubbell, D.S. Cafiso, C. Altenbach, Identifying conformational changes with site-directed spin labeling, Nat. Struct. Biol. 7 (2000) 735–739. H.S. Mchaourab, M.A. Lietzow, K. Hideg, W.L. Hubbell, Motion of spin-labeled side chains in T4 lysozyme. Correlation with protein structure and dynamics, Biochemistry 35 (1996) 7692–7704. S. Onishi, K. Maruyama, S. Ebashi, Calcium-induced conformational changes and mutual interactions of troponin components as studied by spin labeling, J. Biochem. 78 (1975) 73–81. J.A. Spudich, S. Watt, The regulation of rabbit skeletal muscle contraction. I. Biochemical studies of the interaction of the tropomyosin–troponin complex with actin and the proteolytic fragments of myosin, J. Biol. Chem. 246 (1971) 4866–4871.
468
T. Aihara et al. / Biochemical and Biophysical Research Communications 340 (2006) 462–468
[30] S. Ebashi, A. Kodama, F. Ebashi, Troponin. I. Preparation and physiological function, J. Biochem. (Tokyo) 64 (1968) 465– 477. [31] P.C.S. Chong, R.S. Hodges, Proximity of sulfhydryl groups to the sites of interaction between components of the troponin complex from rabbit skeletal muscle, J. Biol. Chem. 257 (1982) 2549–2555. [32] T. Ojima, K. Nishita, Separation of Akazara scallop and rabbit troponin components by a single-step chromatography on CMToyopearl, J. Biochem. (Tokyo) 104 (1988) 9–11. [33] N. Matsuo, Y. Nagata, J. Nakamura, T. Yamamoto, Coupling of calcium transport with ATP hydrolysis in scallop sarcoplasmic reticulum, J. Biochem. (Tokyo) 131 (2002) 375–381.
[34] A. Redfield, The theory of relaxation processes, Adv. Magn. Reson. 1 (1965) 1. [35] D.E. Budil, S. Lee, S. Saxena, J.H. Freed, Non-linear least squares analysis of slow-motion EPR spectra in one and two dimensions using a modified Levenberg–Marquardt algorithm, J. Magn. Reson. 120 (1996) 155–189. [36] H.C. Li, K. Hideg, P.G. Fajer, The mobility of troponin C and troponin I in muscle, J. Mol. Recognit. 10 (1997) 194–201. [37] K. Murakami, F. Yumoto, S.Y. Ohki, T. Yasunaga, M. Tanokura, T. Wakabayashi, Structural basis for Ca2+-regulated muscle relaxation at interaction sites of troponin with actin and tropomyosin, J. Mol. Biol. 352 (2005) 178–201.