FASEB J. Uncover Your Biological Pathway
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Published as doi: 10.1096/fj.07-8630com.
 (The FASEB Journal. 2007;21:3974-3985.)
© 2007 FASEB
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
fj.07-8630comv1
21/14/3974    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Szczesna-Cordary, D.
Right arrow Articles by Lopaschuk, G. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Szczesna-Cordary, D.
Right arrow Articles by Lopaschuk, G. D.

Myosin regulatory light chain E22K mutation results in decreased cardiac intracellular calcium and force transients

Danuta Szczesna-Cordary*,1, Michelle Jones*, Jeffrey R. Moore{ddagger}, James Watt{ddagger}, W. Glenn L. Kerrick{dagger}, Yuanyuan Xu{dagger}, Ying Wang{dagger}, Cory Wagg§ and Gary D. Lopaschuk§

Departments of
* Molecular and Cellular Pharmacology,

{dagger} Physiology and Biophysics, University of Miami, Miller School of Medicine, Miami, Florida, USA;

{ddagger} Department of Physiology and Biophysics, Boston University School of Medicine, Boston, Massachusetts, USA; and

§ Cardiovascular Research Group, University of Alberta, Edmonton, Canada

1Correspondence: University of Miami School of Medicine, Department of Molecular & Cellular Pharmacology (R-189), P.O. Box 016189, 1600 NW 10th Ave, Rm. 6113, Miami, FL 33101, USA. E-mail: dszczesna{at}med.miami.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The glutamic acid to lysine mutation at the 22nd amino acid residue (E22K) in the human cardiac myosin regulatory light chain (RLC) gene causes familial hypertrophic cardiomyopathy (FHC) with a phenotype of midventricular obstruction and septal hypertrophy. Our recent histopathology results have shown that the hearts of transgenic E22K mice (Tg-E22K) resemble those of human patients, demonstrating enlarged interventricular septa and papillary muscles. In this study, we show no effect of the E22K mutation on the kinetics of mutated myosin in its ATP-powered interaction with fluorescently labeled single actin filaments compared to nontransgenic or transgenic wild-type (Tg-WT) control mice. Likewise, no change in cross-bridge dissociation rates (gapp) was observed in freshly skinned papillary muscle fibers. In contrast, maximal force and ATPase were decreased ~20% in Tg-E22K skinned papillary muscle fibers and intracellular [Ca2+] and force transients were significantly decreased in intact papillary muscle fibers from Tg-E22K compared to Tg-WT mice. Moreover, energy metabolism measured in isolated working Tg-E22K mouse hearts perfused under conditions of physiologically relevant levels of metabolic demand was similar in Tg-E22K and control hearts before and after 20 min of no-flow ischemia. Our results suggest that the pathological response observed in the E22K myocardium might be triggered by mutation induced changes in the properties of the RLC Ca2+-Mg2+ site, the state of the Ca2+/Mg2+ occupancy and consequently the Ca2+ buffering ability of the RLC. By decreasing the affinity of the RLC for Ca2+, the E22K mutation most likely promotes a Mg2+-saturated RLC producing less force and ATPase than the Ca2+-saturated RLC of WT fibers. Decreased Ca2+ binding may also lead to faster Ca2+ dissociation kinetics in Tg-E22K intact fibers resulting in decreased duration and amplitude of [Ca2+] and force transients. These changes when placed in vivo would result in higher workloads and consequently cardiac hypertrophy.—Szczesna-Cordary, D., Jones, M., Moore, J. R., Watt, J., Kerrick, W. G. L., Xu, Y., Wang, Y., Wagg, C., Lopaschuk, G. D. Myosin regulatory light chain E22K mutation results in decreased cardiac intracellular calcium and force transients.


Key Words: transgenic mice • Ca2+/Mg2+ binding to RLC • cardiac hypertrophy • in vitro motility assays • intact and skinned muscle fibers • energy metabolism


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
DESPITE ADVANCED MEDICAL interventions, cardiovascular diseases remain a major cause of mortality worldwide. Familial hypertrophic cardiomyopathy (FHC) is one of the pathological compensatory manifestations found in the heart resulting from its inability to adequately pump blood, thus leading to premature fatigue, dyspnea, hypertrophy, and/or cardiac failure (1 , 2) . FHC is an autosomal dominant disease originating from mutations in genes that encode for the major contractile proteins of the heart, including the ventricular regulatory light chain (RLC) of myosin (3 4 5 6 7 8 9) . FHC is characterized by ventricular and septal hypertrophy, myofibrillar disarray, abnormal ECG findings, and sudden cardiac death (10 11 12) . The glutamic acid to lysine mutation at residue 22 (E22K) was one of the first RLC mutations found to be associated with FHC, and its phenotype showed massive hypertrophy of the cardiac papillary muscles and adjacent ventricular tissue (3) . The later diagnostic study of Kabaeva et al. (6 , 13) found this E22K mutation to cause moderate septal hypertrophy, a late onset of clinical manifestation, and benign disease course and prognosis.

To investigate the functional consequences of the E22K mutation located in the Ca2+ binding EF hand motif of the RLC (Fig. 1 ), we generated transgenic mice over-expressing the E22K mutant of human ventricular RLC (14) . Histologically, the hearts of these animals resemble those of human patients demonstrating midventricular obstruction due to septal and papillary muscle hypertrophy (3 , 6) . Longitudinal sections of hematoxylin & eosin stained whole hearts of 13-month-old Tg-E22K mice showed enlarged interventricular septa and papillary muscles compared to nontransgenic (NTg) mice or mice over-expressing the wild-type human cardiac RLC (Tg-WT; ref. 14 ). E22K mutant-reconstituted skinned porcine muscle fibers (15) or glycerinated E22K transgenic papillary muscle fibers (14) showed a slight increase in the Ca2+ sensitivity of force and myofibrillar ATPase activity compared to either bacterially expressed WT-reconstituted or transgenic muscle preparations. This indicated that the E22K mutation may cause structural perturbations in RLC that may affect the properties of the RLC Ca2+-Mg2+ site and its ability to bind Ca2+ and/or Mg2+. These myofilament and RLC-mediated alterations in Ca2+ signaling might trigger abnormal heart function, initiating a series of events that eventually lead to cardiac hypertrophy and/or heart failure.


Figure 1
View larger version (46K):
[in this window]
[in a new window]

  		Figure 1. Schematic representation of E22K mutation (labeled in green) in myosin regulatory light chain (NCBI accession number: 2MYS). Heavy chain of myosin is labeled in blue, essential light chain (ELC) in yellow, and RLC in magenta.

Using transgenic E22K cardiac muscle myofibrils and confocal microscopy, Dumka et al. (16) have recently shown that the E22K mutation has negligible influence on the time of cross-bridge dissociation, rebinding and ADP dissociation. The difference in dissociation time was found to be not statistically significant with 60% of myosin molecules carrying the mutation (Tg-E22K line 2) and marginally significant with 90% mutant myosin (Tg-E22K line 4). Likewise, no effect was observed on the rate of cross-bridge rebinding to thin filaments or on the time of ADP dissociation from the myosin active site (16) .

In this study, we further investigated the functional consequences of the E22K mutation using in vitro motility assays, freshly skinned and intact papillary muscle fibers, and isolated working hearts. Consistent with the results by Dumka et al. (16) , we demonstrate that the E22K mutation has no effect on the kinetics of the interaction of myosin isolated from transgenic hearts with single actin filaments in an in vitro motility assay. Similarly, no change in cross-bridge dissociation rate (gapp) was observed in the freshly skinned papillary muscle fibers. In contrast, an ~20% decrease in maximal force and ATPase was observed in Tg-E22K skinned papillary muscle fibers compared to control Tg-WT fibers. Furthermore, simultaneous intracellular [Ca2+] and force transients in electrically stimulated intact muscle fibers were significantly decreased in Tg-E22K compared to Tg-WT mice. The current study also shows no E22K-induced changes in energy metabolism measured in isolated working transgenic mouse hearts perfused under conditions of physiologically relevant levels of metabolic demand. After 20 min of no flow ischemia, the E22K hearts demonstrated a slightly better recovery compared to WT hearts (17) .


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
All animal studies were conducted in accordance with institutional guidelines. Transgenic mouse models expressing the myc-tagged WT or two lines (L2 and L4) of the E22K mutant of human cardiac RLC generated previously (14 , 16) were utilized in the in vitro, skinned and intact fiber, and isolated perfused heart experiments. Transgenic protein expression levels were myc-WT ~25%, myc-E22K L2 ~65%, and myc-E22K L4 ~87% (14) . For all experiments, the animals were matched by age and gender and the results of transgenic myc-E22K (depicted as Tg-E22K) mouse preparations were compared to nontransgenic (NTg) and/or transgenic myc-WT (depicted as Tg-WT) control preparations. Where no differences between two lines (L2 and L4) of E22K transgenic muscle preparations were found (P>0.05), the results were combined and averaged.

RNA extraction and RT-PCR
Mouse ventricles were rapidly harvested from ~8-mo-old NTg, Tg-WT, Tg-E22K-L2, and Tg-E22K-L4 mice; submerged immediately in an appropriate volume of ice-cold RNAlater reagent (Qiagen, Valencia, CA,USA); and stored frozen at –80°C until needed. Total RNA was isolated using a RNeasy Fibrous Tissue Kit (Qiagen). First strand cDNA was synthesized using 4 µg of total RNA and 1 µM oligo-dT primer using Omniscript reverse transcriptase according to the manufacturer’s protocol (Qiagen). The PCR reaction was performed using 1/10 volume of the RT reaction using sets (sense and antisense) of specific primers designed to amplify the mouse cardiac SERCA 2 (NCBI accession #NM_009722) and phospholamban (PLB; NCBI accession #NM_023129) genes (Table 1 ). Concurrently, a set of primers designed to amplify the mouse cardiac {alpha}-actin gene (NCBI accession #NM_009608) was used as a loading control in each PCR reaction. Final concentrations of SERCA 2 and PLB primers were 0.8 µM, whereas that of mouse {alpha}-actin 0.1 µM. The PCR reactions were performed using TaqDNA polymerase (Invitrogen) in a final volume of 25 µl. The PCR products were electrophoresed on a 1% agarose gel and visualized using ethidium bromide staining. Reactions were documented using a Bio-Rad Gel Imaging System (Gel Doc XR, Bio-Rad Laboratories, Hercules, CA, USA) and Image Quant Software (18) .


View this table:
[in this window]
[in a new window]

  		Table 1. Primers used for analysis of expression pattern of mouse cardiac SERCA 2 and phospholamban genes. NCBI accession numbers: SERCA 2 #NM_009722, PLB #NM_023129 and {alpha}-actin #NM_009608.

Isolation and purification of transgenic myosin
Myosin from ~8 month-old NTg and Tg mouse ventricles was prepared according to the protocol of Svensson et al. (19) . Briefly, ventricles were first minced by hand and then homogenized in an ice-cold Guba Straub type buffer [300 mM NaCl, 100 mM NaH2PO4, 50 mM Na2HPO4, pH 6.5 and 10 mM Na4P2O7, 1 mM MgCl2, 10 mM EDTA, 1 mM DTT, 0.1% NaN3, protease inhibitor cocktail (Sigma, St. Louis. MO, USA)] for 2 min at 20 Hz in a Mixer mill MM300. The homogenate was then incubated on ice for 40 min before centrifugation at 200,000 g for 1 h. The supernatant was then diluted 1:60 v/v with ice-cold 2 mM DTT and incubated on ice 30 min with gentle stirring and left to stand for another 30 min. The samples were then centrifuged at 30,000 g for 10 min, and the pellet was dissolved in a buffer containing 0.6 M KCl, 10 mM MOPS, pH 7.0, 5 mM DTT, 0.1% NaN3, protease inhibitor cocktail. Samples were diluted 1:1 with glycerol, mixed gently and stored at –20°C up to 4–7 days.

In vitro motility assays
The in vitro motility assays were performed as described previously (20) , with some subtle modifications. Flow cells were constructed by forming a channel between nitrocellulose coated coverslips and a standard glass slide with 3M double stick tape (100 µm thickness). Myosin was introduced into the chamber in high salt buffer (300 mM KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, and 1 mM DTT) and allowed to bind to the nitrocellulose surface for 2 min. Sixty microliters of 0.5 mg/ml bovine serum albumin (BSA) in high salt buffer were then passed through the chamber to remove any unbound myosin and to block the remaining surface sites in order to avoid non-specific attachment of actin filaments. After blocking, 60 µl of low salt buffer (25 mM KCl, 25 mM imidazole, 1 mM EGTA, 4 mM MgCl2, and 1 mM DTT) was passed through the chamber to remove any unbound BSA. Tetramethylrhodamine isothiocyanate (TRITC)-labeled actin filaments (~5 nM) in low salt buffer were added and allowed to bind to the myosin in the absence of ATP. Unbound actin was removed by washing with low salt buffer, and movement was initiated by the addition of low salt buffer with 1 mM ATP, scavenger (glucose oxidase, catalase, and dextrose, and 0.5% methylcellulose added.

Filament movement was observed at 25°C with an ICCD camera model IC200 (PTI, Birmingham, NJ, USA). Video sequences were captured using Scion image and an AG-5 image grabber (Scion Corp., Frederick, MD, USA). The average velocity for a given filament was determined from the distance traveled by the filament between 10–12 consecutive video images taken at 1 s intervals using Retrac, the freeware program written by Dr. Nick Carter. Fifteen to twenty five filaments from each video segment were averaged, and at least two video segments were obtained per flow cell.

Estimating myosin duty cycle
The duty cycle of the mutant myosin molecules was estimated by the method of Uyeda et al. (21) . Briefly, actin filaments were translocated at maximum velocity (Vmax) if at least one head was interacting with the filament at all times. The probability of this occurring is related to the number of heads available to interact with actin, N, and the fraction of its cycle time that the myosin molecule remains bound, f (i.e., duty cycle). To determine the number of myosin heads available to interact with an actin filament, we assumed that a myosin molecule 10 nm away was capable of reaching out and attaching to a 5 nm wide actin filament. Therefore, for any length of actin filament, a 25 nm wide band of myosin molecules was assumed to be available to interact with the actin filament. The number of molecules within the band was determined by the surface density estimates ({rho}; ref. 20 , 22 ) and the contact area (A) for an average filament length of 3.1 ± 1.6 µm (N={rho}A; ref. 20 , 23 ).

Assuming that the motors act asynchronously, the probability that at least one out of N heads is interacting with actin at all times is given by 1-(1-f){rho}A. Consequently, at low motor surface densities the velocity will decrease as a function of f and surface density according to the equation:

Formula 1(1)
where Vobs is the observed velocity of the actin filament (21) . To estimate the duty cycle, we measured actin filament velocity as a function of myosin loading concentration (i.e., the number of heads available to interact with actin, see above). Plots of velocity as a function of the number of heads available to interact with a particular filament were then fit by nonlinear regression analysis (SigmaPlot 2000; SPSS Inc., Chicago, IL, USA) to Eq. 1 with the duty cycle, f, as the fit parameter.

Fiber studies
Apparatus
Intact and skinned muscle fiber studies were performed utilizing 7 ± 1-month-old mice matched by gender in the Guth Muscle Research System (Scientific Instruments, Heidelberg, Germany), as described in detail in our previous work (24 , 25) . Briefly, the mechanical parts of the apparatus consisted of a force transducer for measuring force. The optics consisted of a microscope photometer unit for monitoring emission light from the muscle fiber. The light was focused by an Olympus Quartz condenser onto the muscle preparation after passing through filters appropriate for performing [Ca2+] transient and ATPase measurements.

Simultaneous ATPase and force measurements in skinned muscle fibers
After euthanasia, the hearts of mice were excised and strips of mouse papillary muscle fibers (diameter: 60–70 µm) were dissected in ice-cold relaxing solution (85 mM K+, 2 mM MgATP2–, 1 mM Mg2+, 7 mM EGTA, pH 7.0, and propionate as the major anion). The fibers were treated with 1% Triton X-100 for 30 min and processed immediately without glycerination (14) . The skinned fiber was then mounted in the Guth Muscle Research System, and the sarcomere length was adjusted to 2.2 µm by a laser diffraction pattern. The cross-sectional area was calculated based on measurement of the fiber width by microscope and the assumption that the fiber is circular in diameter. The ATPase rate was measured using the NADH fluorescence method (26) . The regeneration of ATP from ADP and phospho(enol) pyruvate (PEP) by the enzyme pyruvate kinase (PK) is coupled to the oxidation of NADH (fluorescent) to NAD (nonfluorescent) by lactate dehydrogenase (LDH). The fiber was subjected to an increasing Ca2+ gradient (a slow, uniform, stepwise increase in Ca2+ concentrations) by using a gradient maker to mix the relaxing and contracting solutions (relaxing: pCa 9.0; contracting: pCa 3.4; ref. 24 ). Both solutions contained 85 mM K+, 2 mM MgATP2–, 1 mM Mg2+, 7 mM EGTA, 10–3.4 or 10–9 M Ca2+, 5 mM PEP, 100 U/ml PK, 0.4 mM NADH, 140 U/ml LDH, ionic strength 150mM, pH 7.0, and propionate as the major anion. A computer-controlled pump replenished the solution in the cuvette every 20 s with a solution of a continuously increasing Ca2+ gradient. Fresh, unoxidized NADH solution was introduced into the cuvette every 20 s. The decrease in NADH concentration was determined by a decrease in the fluorescence signal detected at 450 nm. The slope of the linear decrease in NADH concentration was used to calculate the ATPase rate. The Ca2+ concentration gradient was calibrated by use of the fluorescent Ca2+ indicator calcium green-2 (Molecular Probes, Eugen, OR, USA). A detailed description of the calibration is described by Allen et al. (27) . Force development was monitored simultaneously with ATPase measurements utilizing the force transducer of the Guth apparatus.

Force transient and [Ca2+] transient measurements in intact muscle fibers
Intact mouse papillary muscles (1.0–2.0x0.2–0.3 mm) were dissected free in ice-cold Krebs-Henseleit solution (119 mM NaCl, 4.6 mM KCl, 1.8 mM CaCl2, 1.2 mM MgSO4, 25 mM NaHCO3, 1.2 mM KH2PO4, and 11 mM glucose) containing 30 mM 2,3-butanedione monoxime (BDM; Sigma), saturated with 95% O2 and 5% CO2. The muscle was then mounted in the Guth muscle research apparatus, its length was adjusted until the maximum active twitch force was obtained, and then it was stimulated at 1.0Hz. Afterward, the muscle was loaded with 5 µM Fura-2-AM (Molecular Probes) in an oxygenated Krebs-Henseleit solution containing 0.5% cremophore for 1 h to increase the solubility of Fura-2 in the bathing solution. The Fura-2 fluorescence signals corresponding to the 340- and 380-nm illumination of the preparation were sampled by a signal sorter and recorded by a computer. The time resolution of the 340 nm to 380 nm fluorescence ratio (340/380) measurements was 4 milliseconds. Background fluorescence from 340 and 380 nm excitations of unloaded preparations was subtracted before the fluorescence ratio 340/380 was calculated and plotted along with force data. Although the off-rate of Ca2+ from Fura-2 is fast enough to measure the time course of [Ca2+] transients in intact papillary muscle fibers, the multiple in vivo binding sites for this hydrophobic Ca2+ indicator makes the determination of the absolute cytosolic [Ca2+] inaccurate (28) . Therefore, the 340/380 fluorescence ratios were used to calculate relative changes in the intracellular [Ca2+] transients.

Isolated mouse working heart perfusion (29)
Transgenic mice were deeply anesthetized with pentobarbital sodium (5–10 mg ip) and hearts were excised and placed in ice-cold Krebs-Henseleit bicarbonate (KHB) solution (118 mM NaCl, 25 mM NaHCO3, 4.7 mM KCl, 1.2 mM KH2PO4, 2.5 mM CaCl2, and 5.0 mM glucose, pH 7.4). Hearts were cannulated first via the aorta with a 15-gauge plastic cannula and perfused retrogradely by the Langendorff method with KHB solution (gassed with 95% O2-5% CO2). During this time, the left atrium was cannulated through a pulmonary opening with a 16-gauge steel cannula. Once the cannulation for the working heart perfusions was complete, the Langendorff line was closed and the left atrial and aortic lines were opened and perfused with KHB solution containing 5.0 mM glucose, 1.2 mM palmitate bound to 3% fatty acid-free BSA at a preload pressure of 11.5 mmHg and an afterload pressure of 50 mmHg. The perfusion protocol included a 30 min aerobic perfusion of spontaneously beating hearts followed by a 20 min global ischemia and then a 40 min reperfusion with oxygenated KHB solution. The spontaneous recovery of heart function was monitored. To determine palmitate oxidation, glucose oxidation, and glycolytic rates, trace amounts of [9,10-3H] palmitate (0.1 µCi/ml), [U-14C] glucose (0.1 µCi/ml) or [5-3H] glucose (0.1 µCi/ml) were used, respectively. Functional measurements were acquired for 10 s every 10 min with the MP100 system from AcqKnowledge (BioPac Systems, Santa Barbara, CA, USA). Cardiac output and aortic flows were measured with inline flow probes (Transonic Systems, Ithaca, NY, USA) placed in the left atrial line and the aortic afterload line, respectively. Coronary flow was calculated as the difference between cardiac output and aortic flows. Heart rate and pressures were measured with a pressure transducer (TSD104A, BioPac Systems) placed in the aortic line at the level of the heart. Cardiac work was calculated as the product of peak systolic pressure and cardiac output. Stroke volume was calculated as the cardiac output divided by the heart rate, whereas stroke work was calculated as the stroke volume times the peak systolic pressure. Cardiac power was calculated as the product of developed pressure and cardiac output. A conversion factor of 1.33 x 10–4 was used to convert cardiac power values from millimeters of mercury per milliliter to joules (30) .

Statistical analysis
Data are expressed as the average of n experiments ± SE. Statistically significant differences were determined utilizing an unpaired Student’s t test with significance defined as P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
In vitro motility assays
To test whether the E22K mutation alters the movement of the RLC binding domain of the myosin head during cardiac muscle contraction (20 , 31 32 33) , we examined the actin filament velocity for E22K transgenic myosin in an in vitro motility assay. Actin filament velocities were determined for at least two independent mouse heart preparations from each line. Myosin purified from Tg-WT mice exhibited velocities that were indistinguishable from that purified from NTg mice [1.3±0.04 µm/s vs. 1.4±0.03; P=0.23 for Tg-WT (n=6) and NTg (n=4), respectively]. Tg-E22K L2 (n=2) and Tg-E22K L4 (n=10) were not significantly different (P=0.72) and were therefore pooled. When compared to Tg-WT control, the Tg-E22K myosin also translocated actin filaments at similar velocities (P=0.67; Fig. 2 ). In summary, the E22K mutation did not affect the mechanical properties of the myosin lever arm and did not introduce compliance to the myosin neck region.


Figure 2
View larger version (12K):
[in this window]
[in a new window]

  		Figure 2. In vitro motility assay of Tg-E22K and control (NTg and Tg-WT) mice. Bar graph indicates velocities (µm/s) as mean ± SE. Myosin from Tg-WT mice (n=6) exhibited velocities that were indistinguishable from that of NTg mice (n=4; P=0.23). Tg-E22K L2 (n=2) and Tg-E22K L4 (n=10) were not significantly different (P=0.72) and were therefore pooled. No statistically significant differences in velocities of Tg-E22K mice and control NTg and Tg-WT mice were observed (P=0.67).

To further determine if the E22K mutation is associated with alterations in actin-myosin kinetic transitions, each myosin was analyzed for its ability to move actin in the in vitro motility assay over a range of myosin concentrations (see Materials and Methods for experimental detail). The results from these experiments (Fig. 3 ) show that actin filament velocity was independent of myosin concentration over a broad range (30–150 µg/ml myosin). However, as the concentration of myosin loaded onto the motility surface was lowered below 30 µg/ml, the number of filaments moving on the surface was reduced, and the actin filament motion became more erratic and the average velocity decreased. The rate at which the velocity decreases provides a measure of the duty cycle, which is the fraction of the total ATPase cycle that the myosin molecule remains attached to actin (see Materials and Methods for details). There were no significant differences in duty cycle for any of the myosin preparations tested (Fig. 3) . Also, similar to what was predicted from Fig. 2 , fits of the data in Fig. 3 to Eqn. 1 point out that myosin preparations purified from Tg-E22K mice show no deficits in maximal actin filament velocity (Vmax=1.2±0.06, 1.27±0.04, and 1.24±0.07 for NTg, Tg-WT, and Tg-E22K, respectively). Overall, the in vitro motility assay data indicate that, although in a critical region for myosin motion generation, the E22K mutation does not affect myosin mechanics or biochemistry under the lightly loaded conditions of the in vitro motility assay.


Figure 3
View larger version (8K):
[in this window]
[in a new window]

  		Figure 3. The dependence of actin filament velocity on the number of interacting myosin molecules. Average velocity (µm/s± SE) is plotted as a function of myosin motor surface density. A, B, C) are NTg, Tg-WT, and Tg-E22K respectively. Note dependence of actin filament velocity on the number of interacting heads. Myosin motility data were best fit by theoretical curves (Eqn. 1) representing duty cycles of f = 0.034 (3.4%), 0.034 (3.4%), and 0.027 (2.7%) for NTg, Tg-WT and Tg-E22K respectively (solid lines). The data were obtained for 2 independent myosin preparations from NTg, Tg-WT and 4 myosin preparations from Tg-E22K mice. 25 filaments from 2–4 movies were analyzed for each concentration of myosin (each point on the plot).

Skinned fiber studies
Simultaneous ATPase-pCa and force-pCa measurements (Fig. 4 and Fig. 5 A, B) were carried out under isometric conditions in freshly skinned papillary muscle fibers from Tg-E22K (L2 and L4) and Tg-WT mice. All fiber experiments were performed on 7 ± 1.0-month-old mice matched by gender. The results showed no differences between two transgenic E22K lines (L2 and L4) and were therefore analyzed together. Figure 4 demonstrates the relationship between the maximum force and a maximal level of ATPase activity. Skinned fibers from Tg-E22K mice (n=16 fibers) showed a significant 20% decrease in both maximum ATPase (Fig. 4 left panel) and maximum force (Fig. 4 , right panel) compared to Tg-WT controls (P<0.01, n=19 fibers). Figure 5A shows the Ca2+ sensitivity of simultaneously monitored force and ATPase in these freshly skinned papillary muscle fibers. Contrary to what we previously reported for glycerinated skinned muscle fibers (14) , no significant differences in the pCa50 of force/ATPase-pCa relationships were observed between Tg-E22K and Tg-WT in freshly skinned papillary muscle fibers used in this study (Fig. 5A ). This suggests that freshly skinned preparations somewhat differ from skinned glycerinated fibers. Figure 5B demonstrates the ratio of the actomyosin ATPase activity to force as a function of the state of activation (normalized force). Consistent with the in vitro motility results, the energy cost or the rate of dissociation of myosin cross-bridges (ATPase/force{approx}gapp) was not altered in Tg-E22K compared to Tg-WT fibers. In addition, as shown in Fig. 5A , the shape of the force-pCa and ATPase-pCa curves did not change, which suggests that the rate of association of force generating myosin cross-bridges (fapp) is also not changed by the E22K mutation of RLC.


Figure 4
View larger version (10K):
[in this window]
[in a new window]

  		Figure 4. Maximal ATPase (left panel) and force (right panel) measurements performed simultaneously in freshly skinned papillary muscle fibers from Tg-E22K and Tg-WT mice under isometric conditions. Note a significant 20% decrease in maximum ATPase (P=0.0058) and maximum force (P=0.00004) in skinned fibers from Tg-E22K mice (n=16 fibers) compared to Tg-WT mice (n=19 fibers).


Figure 5
View larger version (13K):
[in this window]
[in a new window]

  		Figure 5. ATPase/force-pCa relationships (A) and ratio of ATPase/force {cong} gapp (in s–1 per myosin head/ x 105 N/m2 (B) assessed simultaneously in freshly skinned papillary muscle fibers from Tg-E22K (n=16) and Tg-WT (n=19) mice.

Intact fiber studies
Intact papillary muscle fibers from Tg-E22K and Tg-WT mice were stimulated at 1 Hz, and force and fluorescence originating from Fura-2 were monitored simultaneously. As in skinned fiber experiments, the results from intact fiber measurements showed no differences between two transgenic E22K lines and were therefore combined. Normalized force (Fig. 6 A) and [Ca2+] transients (Fig. 6B ) for Tg-E22K and Tg-WT intact fibers were plotted as a function of time in milliseconds (ms). As pointed out by Baylor and Hollingworth (28) , Fura-2 cannot be calibrated to measure accurately intracellular Ca2+ due to its hydrophobic interactions with intracellular proteins that most likely alter the Kd of Fura-2 for Ca2+. Since the ratio of bound to unbound Fura-2 to protein is not known the Kd of Fura-2 for Ca2+ cannot be accurately determined. Therefore, we have not attempted to measure intracellular Ca2+ concentrations but the time course of [Ca2+] transients (28) .


Figure 6
View larger version (10K):
[in this window]
[in a new window]

  		Figure 6. Normalized force (A, C) and [Ca2+] (B, D) transients in electrically stimulated intact papillary muscle fibers from Tg-E22K and Tg-WT mice. Note significant differences in force (P<0.03) and [Ca2+] (P<0.03) transients between E22K (n=5) and WT (n=4) mouse fibers.

Table 2 summarizes the t50 and t10 values characterizing the time (in ms) from the peak to 50 and 10%, respectively, of force and [Ca2+] in relaxation. The E22K mutation caused significant functional changes in intact papillary muscle fibers compared to Tg-WT muscle fibers. Force (Fig. 6A ) and [Ca2+] (Fig. 6B ) transients were significantly shortened in Tg-E22K mice and the t50 and t10 values of force transients were respectively 1.34- and 1.54-fold lower in Tg-E22K papillary muscles compared to those of Tg-WT (Table 2) . Likewise, the duration of [Ca2+] transients in Tg-E22K papillary muscles was ~20% shorter than in Tg-WT, judging by the t50 and t10 values in relaxation (Table 2 ; Fig. 6B ). To determine the minimum relative change in peak force and intracellular Ca2+ between Tg-E22K and Tg-WT muscle fibers, the normalized force and [Ca2+] transients curves were adjusted so that the rise time of the force and [Ca2+] transients coincided. Because the skinned fiber data indicated that there was no change in Ca2+ sensitivity or cross-bridge kinetics, the rise time of the higher magnitude Tg-WT should be at least as fast as the lower magnitude Tg-E22K force transient. Additionally, the rise time of the higher magnitude Tg-WT [Ca2+] transient should be at least as fast as the lower magnitude Tg-E22K [Ca2+] transient. These adjustments to the data from Fig. 6A, B are shown in Fig. 6C, D . The magnitude and duration of the force and [Ca2+] transients in Tg-E22K muscles were decreased compared to Tg-WT muscle fibers (Fig. 6C, D ). Thus, the functional consequence of the E22K mutation in RLC is to decrease the magnitude and duration of the force transient during twitch contraction.


View this table:
[in this window]
[in a new window]

  		Table 2. Force and [Ca2+] transients in intact papillary muscle fibers from Tg-E22K and Tg-WT mice

Expression of Ca2+-handling proteins
Given that the E22K mutation affects intracellular [Ca2+] and force transients, we have examined the expression levels of two cellular calcium handling proteins, the sarcoplasmic reticulum (SR) Ca2+-dependent ATPase (SERCA 2) and its regulatory protein PLB. As shown in Fig. 7 A, B, the levels of mRNA expression for SERCA 2 and PLB were not different among NTg, Tg-WT and Tg-E22K (L2 and L4) mouse extracts, identified with the specific primers designed to amplify the mouse cardiac SERCA 2 and PLB genes (Table 1) . Mouse cardiac {alpha}-actin (identified with the specific primers listed in Table 1 ) was used as an internal loading control. Figure 7C shows the protein expression of the monomeric form of PLB and demonstrates no difference in all examined NTg, Tg-WT, and Tg-E22K ventricular tissue lysates. The upper panel of Fig. 7C demonstrates the expression of mouse cardiac RLC (depicted as RLCendog) and transgenic human ventricular RLC (depicted as myc-RLC) in all samples that were blotted for PLB. The RLC protein expression was identified with polyclonal RLC-specific antibodies produced in our laboratory (14 , 25) . In agreement with what we reported earlier (14) , the amounts of transgene expression were: ~25, ~65, and ~87% in Tg-WT, Tg-E22K L2, and Tg-E22K L4 mouse lines, respectively. These data suggest that the E22K mutation did not affect either mRNA or protein expression of SERCA 2 and PLB.


Figure 7
View larger version (29K):
[in this window]
[in a new window]

  		Figure 7. Analysis of mRNA and protein expression of SR Ca2+ handling genes in NTg, Tg-WT and Tg-E22K mouse ventricular extracts. SERCA 2 mRNA/PCR products (A), PLB (B) mRNA/PCR products, and PLB protein expression (C) in left ventricular mouse extracts from NTg, Tg-WT, and Tg-E22K L2 and Tg-E22K L4 mice. A, B) Lanes 1 and 5 = 100 bp and 1 Kb Plus DNA ladders, respectively; lanes 2, 3, 4 and 5, = NTg, Tg-WT, Tg-E22K L2, and Tg-E22K L4. C) PLB protein expression in NTg (lane 1), Tg-WT (lane 2), Tg-E22K L2 (lane 3) and Tg-E22K L4 (lane 4) mice. Ten micrograms of ventricular extracts (1:1 lysis buffer and Laemmli loading buffer) were electrophoresed on 15% SDS polyacylamide gels, and transferred to nitrocellulose membrane (Bio-Rad, Hercules). Blocking and overnight antibody incubations were done at room temperature. Membranes were blocked 30 min in a 50:50 mix of Tris buffered saline, 0.05% Tween 20: Rockland blocking buffer (T-TO) for Odyssey fluorescence detection. PLB was detected with monoclonal anti-PLB antibody MA3–922 (Affinity BioReagents). The secondary antibody was goat anti-mouse Cy5.5 (Rockland) at a 1:4000 dilution in T-TO. RLC was detected with rabbit polyclonal anti-RLC CT-1 antibody developed in this laboratory (at a 1:2000 dilution). The specific RLC antibody was detected with a 1:4000 dilution of goat anti-rabbit IgG antibody labeled with IR Red 800 fluorescent dye (Rockland). Reaction signal was measured by scanning the blot with the Odyssey Infrared Imager (LI-COR; refs. 14 , 25 ).

Energy metabolism
We also examined the ability of Tg-E22K hearts to perform work under conditions of physiologically relevant levels of metabolic demands (34) and investigated the ability of these hearts to recover after a severe ischemic episode. In addition, glucose and fatty acid oxidation were measured in the isolated working mouse hearts under aerobic conditions and during reperfusion after 20 min of global ischemia (17) . As shown in Fig. 8 A and Table 3 , cardiac work and cardiac power were similar in Tg-E22K mice hearts compared to control Tg-WT or NTg mice during the initial aerobic perfusion period. When reperfused after 20 min of ischemia, hearts from both the Tg-E22K and Tg-WT mice were more susceptible to cardiac injury than hearts from NTg mice (Fig. 8A ). Tg-E22K mice demonstrated a recovery profile that was not significantly different from Tg-WT hearts (P=0.38). No differences in the rates of glucose or palmitate oxidation were observed between Tg-E22K, Tg-WT or NTg mouse hearts, either before or after the ischemic episode (Fig. 7B-D ; Table 3 ). This suggests that the E22K mutation did not have any specific effect on energy metabolic preference or on specific ATP production (Fig. 8 ; Table 3 ).


Figure 8
View larger version (13K):
[in this window]
[in a new window]

  		Figure 8. Energy metabolism in Tg-E22K mice compared to control, Tg-WT and NTg mice. A) Cardiac power in Tg-E22K (n=9), Tg-WT (n=6) and NTg (n=10) mouse hearts. Note slightly better but not statistically significant (P=0.38) recovery in reperfused Tg-E22K mouse hearts compared to reperfused hearts of Tg-WT mice. Glucose oxidation (B), palmitate oxidation (C), and glycolysis rates (D) in isolated working mouse hearts from Tg-E22K, Tg-WT and NTg mice under aerobic conditions and during reperfusion after 20 min of global ischemia.


View this table:
[in this window]
[in a new window]

  		Table 3. Energy metabolism measurements performed in isolated perfused working hearts from Tg-E22K and control mice


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Myosin RLC is one of the sarcomeric proteins of the heart shown to be associated with FHC. The E22K mutation of RLC causes a rare phenotype of FHC characterized by midventricular cardiac obstruction due to massive papillary muscles and adjacent ventricular tissue (3) . Studies by Kabaeva et al. (6 , 13) further revealed an additional phenotype of moderate septal hypertrophy associated with the E22K mutation of RLC. No cases of sudden cardiac death have been reported in any of the E22K patients or their families. To study the mechanism by which the E22K mutation in RLC results in ventricular and/or septal hypertrophy and alters cardiac muscle contraction in humans, we generated transgenic mice over-expressing human cardiac E22K-RLC (14) . The gross morphology of hematoxylin & eosin stained longitudinal sections of Tg-E22K mouse hearts showed that their interventricular septa as well as their papillary muscles were larger than those of Tg-WT or NTg littermates. Thus, previous findings demonstrated that the human phenotype of the E22K-mutated hearts could be recapitulated in transgenic mice (14) .

In this study, we further examined the physiological consequences of this mutation at the single molecule, fiber, and isolated working heart levels. Our goal was to identify the mechanism by which the E22K mutation initiates physiological changes in the myocardium that ultimately lead to ventricular/septal hypertrophy in humans harboring this myosin RLC mutation. A number of hypotheses have been put forward to explain the mechanisms by which sarcomeric protein mutations lead to hypertrophy and/or heart failure in humans. One hypothesis is that myofilament incorporation of mutant proteins depresses contractile function (35) and, as a consequence, the heart tries to compensate (36) . This could be due to the mutation-mediated changes in the kinetics of myosin cross-bridge interaction with actin, increased energy demands, and/or alterations in the Ca2+ homeostasis in the mutated myocardium. In this study, we show that the E22K mutation does not affect the velocity of actin filaments moving along the E22K mutant myosin-coated surface in an in vitro motility assay. These findings agree with original results by Poetter et al. (3) , where the E22K-mutated myosin isolated from cardiac biopsies of affected individuals showed normal actin filament translocation compared to control samples. We conclude that the E22K mutation exerts no effect on the motion of the myosin lever arm during interaction of myosin cross-bridges with actin filaments. In support of this, we observed no change in cross-bridge dissociation rate (gapp) expressed as the ratio of ATPase/force and plotted as a function of the state of activation in our skinned fiber experiments. This suggests that the energy cost is not altered by the E22K mutation of myosin RLC. The results from skinned fiber measurements were fully supported by the experiments showing no significant changes in energy metabolism measured in isolated intact Tg-E22K mouse hearts under conditions of physiologically relevant levels of metabolic demand. The E22K mutation also did not affect cardiac power in aerobic conditions (Table 3) .

The fact that the E22K mutation produced no myofilament kinetic and energetic changes prompted us to investigate the effect of the mutation on overall Ca2+ homeostasis in the myocardium. As found with other members of the superfamily of EF-hand calcium binding proteins, the RLC possess a functional helix-loop-helix Ca2+-Mg2+ binding site whose intracellular concentration is about 2-fold that of the regulatory Ca2+ specific sites of cardiac troponin C (cTnC), the major myofilament Ca2+ buffer. We therefore hypothesized that binding of Ca2+ to RLC is important for intracellular Ca2+ signaling. One can assume that RLC functions as a temporary intracellular buffer for Ca2+ that would work in parallel with the SR Ca2+ pump to sequester Ca2+ and promote relaxation in muscle. By changing the properties of the RLC Ca2+-Mg2+ binding site, the E22K mutation could affect the function of RLC as a delayed Ca2+ buffer. The affinity of Ca2+ binding to recombinant human cardiac RLC was shown to be ~105 M–1 (37) . However, when the RLC is bound to the heavy chain of myosin, the binding increases to ~107 M–1 (38) . As we previously showed, the E22K mutation decreases binding of Ca2+ to RLC by ~17-fold (37) and possibly modifies the specificity of the RLC EF-hand site. One can hypothesize that the E22K mutation makes the Ca2+-Mg2+ binding site of RLC behave like the N-terminal regulatory site of cTnC, which exhibits low affinity and faster Ca2+ dissociation kinetics, thus spawning an additional myofilament Ca2+ buffer. Consequently, a faster Ca2+ reuptake and the shorter duration of [Ca2+] and force transients would be expected, both indicative of enhanced muscle relaxation. Consistent with this hypothesis, our intact muscle fiber experiments (summarized in Fig. 6 and Table 2 ) strongly suggest that the E22K mutation increases the kinetics of Ca2+ reuptake at low intracellular Ca2+ concentrations. In support of this explanation, we have recently reported that another RLC mutation, R58Q, which inactivated RLC for Ca2+ binding, induced an opposite effect to E22K in intact papillary muscle fibers (25) . Prolonged [Ca2+] and force transients were monitored in R58Q preparations suggesting impaired muscle relaxation (25) . Examination of the key cellular Ca2+ reuptake proteins, the SR Ca2+ pump (SERCA 2) and its regulatory protein PLB revealed no effect of the E22K mutation on their mRNA and/or protein expression levels (Fig. 7) . Therefore, the Ca2+-dependent functional changes that we observed in our fiber experiments are essentially caused by the E22K-induced myofilament protein-protein interactions and not by the change in the composition of the SR Ca2+ handling proteins.

Another hypothesis put forward to explain the E22K-mediated effects pertains to the mutation controlled metal occupancy of the Ca2+-Mg2+ binding site of RLC and the mechanism by which Ca2+ or Mg2+ binding to RLC may influence the interaction of myosin with actin and tension generation. Previous studies have shown that an intact Ca2+-Mg2+ binding site of striated, smooth, or scallop myosin RLC is essential for the regulation of the actin-myosin interaction (39 40 41 42) . As mentioned above, the E22K mutation decreases binding of Ca2+ to RLC by ~17-fold (37) and it is likely that even at high intracellular Ca2+ concentrations, the mutation may promote a Mg2+-saturated RLC. In accord with our earlier studies that showed a lower affinity of the Mg2+ over Ca2+ bound heavy meromyosin (HMM) to actin (43) , the Mg2+-saturated RLC in Tg-E22K skinned fibers would produce less maximum force and ATPase than the Ca2+ saturated Tg-WT fibers. Similar results were reported for the skinned skeletal muscle fibers, where ~20% reduction in steady state force was observed when the Ca2+ binding site mutant of RLC (D47A) was exchanged for the endogenous RLC protein (44) . Interestingly, this aspartic acid to alanine mutation at the –Z position of the RLC Ca2+-Mg2+ binding site dramatically decreases Ca2+ binding to RLC (40) , the phenomenon observed in E22K due to FHC (37) . Weaker Ca2+ binding to Tg-E22K vs. Tg-WT intact muscle fibers could also be responsible for its faster dissociation and ultimately decreased duration and amplitude of the [Ca2+] transients observed in Tg-E22K compared to Tg-WT fibers. Therefore, the structural change introduced by the substitution of the conserved glutamic acid with lysine at the amino acid 22 of RLC is most likely communicated to the RLC Ca2+-Mg2+ binding site, located at the residues 37–48 of RLC, and triggers a series of molecular events resulting in impaired Ca2+ binding, decreased force and ATPase, and increased Ca2+ dissociation that would lead to reduced [Ca2+] and force transients in the mutated myocardium. Beyond the structural change brought about by the E to K replacement, the charge change of the E22K mutation from the negatively charged glutamic acid to the positively charged lysine may contribute to the putative physiology of the mutant protein.

One more question to ponder in analyzing the effects of the E22K mutation is whether this FHC-induced structural and charge change in RLC affects the nearby phosphorylation site of RLC, located at serine 15, and the processes that are normally regulated by RLC phosphorylation. In fact, our solution studies demonstrated that this E22K mutation resulted in the mutant protein’s inability to become phosphorylated (37) and this could also take place in vivo. One would anticipate that the lack of RLC phosphorylation contributed to the E22K-mediated functional alterations of the RLC- and Ca2+-dependent intracellular processes.

In summary, in this study we show that the E22K mutation of RLC does not alter cross-bridge kinetics or energy metabolism accompanying cardiac muscle contraction. Instead, the pathological response observed in the E22K mutated myocardium is most likely due to changes in the properties of the RLC Ca2+-Mg2+ site, the state in the Ca2+/Mg2+ occupancy, and consequently the Ca2+ buffering ability of the RLC. Observed functional changes of decreased force/ATPase and reduced [Ca2+] and force transients when placed in vivo would result in a reduced stroke volume and a compensatory increase in heart rate to maintain cardiac output. As a result, the hearts of E22K-mutated patients would be subjected to higher workloads that over a long period of time would lead to cardiac muscle hypertrophy.


   ACKNOWLEDGMENTS
 
We thank Dr. Yingcai Wang for his expertise in analyzing the mRNA gene expression. This work was supported by NIH-HL071778 and AHA Grant-In-Aid 0355384B (to D.S.-C.), NIH-HL077280 and AHA-0435434T (to J. Moore) and a grant from the Canadian Institute for Health Research (to G.D.L.).

Received for publication March 23, 2007. Accepted for publication June 7, 2007.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Lips, D. J., deWindt, L. J., van Kraaij, D. J. W., Doevendans, P. A. (2003) Molecular determinants of myocardial hypertrophy and failure: alternative pathways for beneficial and maladaptive hypertrophy. Eur. Heart J. 24,883-896[Abstract/Free Full Text]
  2. Heineke, J., Molkentin, J. D. (2006) Regulation of cardiac hypertrophy by intracellular signalling pathways. Nat. Rev. Mol. Cell Biol. 7,589-600[CrossRef][Medline]
  3. Poetter, K., Jiang, H., Hassanzadeh, S., Master, S. R., Chang, A., Dalakas, M. C., Rayment, I., Sellers, J. R., Fananapazir, L., Epstein, N. D. (1996) Mutations in either the essential or regulatory light chains of myosin are associated with a rare myopathy in human heart and skeletal muscle. Nat. Genet. 13,63-69[CrossRef][Medline]
  4. Flavigny, J., Richard, P., Isnard, R., Carrier, L., Charron, P., Bonne, G., Forissier, J. F., Desnos, M., Dubourg, O., Komajda, M., et al (1998) Identification of two novel mutations in the ventricular regulatory myosin light chain gene (MYL2) associated with familial and classical forms of hypertrophic cardiomyopathy. J. Mol. Med. 76,208-214[CrossRef][Medline]
  5. Andersen, P. S., Havndrup, O., Bundgaard, H., Moolman-Smook, J. C., Larsen, L. A., Mogensen, J., Brink, P. A., Borglum, A. D., Corfield, V. A., et al (2001) Myosin light chain mutations in familial hypertrophic cardiomyopathy: phenotypic presentation and frequency in Danish and South African populations. J. Med. Genet. 38,E43[CrossRef][Medline]
  6. Kabaeva, Z. T., Perrot, A., Wolter, B., Dietz, R., Cardim, N., Correia, J. M., Schulte, H. D., Aldashev, A. A., Mirrakhimov, M. M., Osterziel, K. J. (2002) Systematic analysis of the regulatory and essential myosin light chain genes: genetic variants and mutations in hypertrophic cardiomyopathy. Eur. J. Hum. Genet. 10,741-748[CrossRef][Medline]
  7. Richard, P., Charron, P., Carrier, L., Ledeuil, C., Cheav, T., Pichereau, C., Benaiche, A., Isnard, R., Dubourg, O., Burban, M., . for the EUROGENE Heart Failure Projectet al (2003) Hypertrophic cardiomyopathy: distribution of disease genes, spectrum of mutations, and implications for a molecular diagnosis Strategy. Circulation 107,2227-2232[Abstract/Free Full Text]
  8. Morner, S., Richard, P., Kazzam, E., Hellman, U., Hainque, B., Schwartz, K., Waldenstrom, A. (2003) Identification of the genotypes causing hypertrophic cardiomyopathy in northern Sweden. J. Mol. Cell. Cardio. 35,841-849[CrossRef][Medline]
  9. Hougs, L., Havndrup, O., Bundgaard, H., Kober, L., Vuust, J., Larsen, L. A., Christiansen, M., Andersen, P. S. (2005) One third of Danish hypertrophic cardiomyopathy patients have mutations in MYH7 rod region. Eur. J. Hum. Genet. 13,161-165[CrossRef][Medline]
  10. Maron, B. J., Olivotto, I., Spirito, P., Casey, S. A., Bellone, P., Gohman, T. E., Graham, K. J., Burton, D. A., Cecchi, F. (2000) Epidemiology of hypertrophic cardiomyopathy-related death: revisited in a large non-referral-based patient population. Circulation 102,858-864[Abstract/Free Full Text]
  11. Maron, B. J. (2002) The young competitive athlete with cardiovascular abnormalities: causes of sudden death, detection by preparticipation screening, and standards for disqualification. Card. Electrophysiol. Rev. 6,100-103[CrossRef][Medline]
  12. Maron, B. J. (2002) Hypertrophic cardiomyopathy: a systematic review. JAMA 287,1308-1320[Abstract/Free Full Text]
  13. Kabaeva, Z. T. (2002) Genetic analysis in hypertrophic cardiomyopathy: missense mutations in the ventricular myosin regulatory light chain gene The Humboldt University of BerlinPh.D. Dissertation
  14. Szczesna-Cordary, D., Guzman, G., Zhao, J., Hernandez, O., Wei, J., Diaz-Perez, Z. (2005) The E22K mutation of myosin RLC that causes familial hypertrophic cardiomyopathy increases calcium sensitivity of force and ATPase in transgenic mice. J. Cell Sci. 118,3675-3683[Abstract/Free Full Text]
  15. Szczesna-Cordary, D., Guzman, G., Ng, S. S., Zhao, J. (2004) Familial hypertrophic cardiomyopathy-linked alterations in Ca2+ binding of human cardiac myosin regulatory light chain affect cardiac muscle contraction. J. Biol. Chem. 279,3535-3542[Abstract/Free Full Text]
  16. Dumka, D., Talent, J., Akopova, I., Guzman, G., Szczesna-Cordary, D., Borejdo, J. (2006) E22K mutation of RLC that causes familial hypertrophic cardiomyopathy in heterozygous mouse myocardium: effect on cross-bridge kinetics. Am. J. Physiol. Heart Circ. Physiol. 291,H2098-H2106[Abstract/Free Full Text]
  17. Wagg, C., Lopaschuk, G. D., Guzman, G., Szczesna-Cordary, D. (2006) Energy metabolism in mutant myosin regulatory light chain transgenic mice. Bioph. J. (Abstracts) ,274a
  18. Wang, Y., Szczesna-Cordary, D., Craig, R., Diaz-Perez, Z., Guzman, G., Miller, T., Potter, J. D. (2007) Fast skeletal muscle regulatory light chain is required for fast and slow skeletal muscle development. FASEB J. 21,2205-2214[Abstract/Free Full Text]
  19. Svensson, C., Morano, I., Arner, A. (1997) In vitro motility assay of atrial and ventricular myosin from pig. J. Cell Biochem. 67,241-247[CrossRef][Medline]
  20. Moore, J. R., Krementsova, E. B., Trybus, K. M., Warshaw, D. M. (2001) Myosin V exhibits a high duty cycle and large unitary displacement. J. Cell Biol. 155,625-635[Abstract/Free Full Text]
  21. Uyeda, T. Q., Abramson, P. D., Spudich, J. A. (1996) The neck region of the myosin motor domain acts as a lever arm to generate movement. Proc. Natl. Acad. Sci. U. S. A. 93,4459-4464[Abstract/Free Full Text]
  22. Harris, D. E., Warshaw, D. M. (1993) Smooth and skeletal muscle myosin both exhibit low duty cycles at zero load in vitro. J. Biol. Chem. 268,14764-14768[Abstract/Free Full Text]
  23. Reck-Peterson, S. L., Tyska, M. J., Novick, P. J., Mooseker, M. S. (2001) The yeast class V myosins, Myo2p and Myo4p, are nonprocessive actin-based motors. J. Cell Biol. 153,1121-1126[Abstract/Free Full Text]
  24. Wang, Y., Xu, Y., Guth, K., Kerrick, W. G. (1999) Troponin C regulates the rate constant for the dissociation of force- generating myosin cross-bridges in cardiac muscle. J. Muscle. Res. Cell Motil. 20,645-653[CrossRef][Medline]
  25. Wang, Y., Xu, Y., Kerrick, W. G. L., Wang, Y., Guzman, G., Diaz-Perez, Z., Szczesna-Cordary, D. (2006) Prolonged Ca2+ and force transients in myosin RLC transgenic mouse fibers expressing malignant and benign FHC mutations. J. Mol. Biol. 361,286-299[CrossRef][Medline]
  26. Guth, K., Wojciechowski, R. (1986) Perfusion cuvette for the simultaneous measurement of mechanical, optical and energetic parameters of skinned muscle fibres. Plügers Arch. 407,552-557[CrossRef][Medline]
  27. Allen, K., Xu, Y. Y., Kerrick, W. G. (2000) Ca2+ measurements in skinned cardiac fibers: effects of Mg2+ on Ca2+ activation of force and fiber ATPase. J. Appl. Physiol. 88,180-185[Abstract/Free Full Text]
  28. Baylor, S. M., Hollingworth, S. (2000) Measurement and interpretation of cytoplasmic [Ca2+] signals from calcium-indicator dyes. News Physiol. Sci. 15,19-26[Abstract/Free Full Text]
  29. Sambandam, N., Morabito, D., Wagg, C., Finck, B. N., Kelly, D. P., Lopaschuk, G. D. (2006) Chronic activation of PPAR{alpha} is detrimental to cardiac recovery after ischemia. Am. J. Physiol. Heart Circ. Physiol. 290,H87-H95[Abstract/Free Full Text]
  30. Belke, D. D., Larsen, T. S., Lopaschuk, G. D., Severson, D. L. (1999) Glucose and fatty acid metabolism in the isolated working mouse heart. Am. J. Physiol. Comp. Reg. Physiol. 277,R1210-R1217
  31. Baker, J. E., Brust-Mascher, I., Ramachandran, S., LaConte, L. E., Thomas, D. D. (1998) A large and distinct rotation of the myosin light chain domain occurs upon muscle contraction. Proc. Natl. Acad. Sci. U. S. A. 95,2944-2949[Abstract/Free Full Text]
  32. Moore, J. R., Krementsova, E. B., Trybus, K. M., Warshaw, D. M. (2004) Does the myosin V neck region act as a lever?. J. Muscle. Res. Cell Motil. 25,29-35[CrossRef][Medline]
  33. Warshaw, D. M., Guilford, W. H., Freyzon, Y., Krementsova, E., Palmiter, K. A., Tyska, M. J., Baker, J. E., Trybus, K. M. (2000) The light chain binding domain of expressed smooth muscle heavy meromyosin acts as a mechanical lever. J. Biol. Chem. 275,37167-37172[Abstract/Free Full Text]
  34. Campbell, F. M., Kozak, R., Wagner, A., Altarejos, J. Y., Dyck, J. R., Belke, D. D., Severson, D. L., Kelly, D. P., Lopaschuk, G. D. (2002) A role for peroxisome proliferator-activated receptor alpha (PPARalpha) in the control of cardiac malonyl-CoA levels: reduced fatty acid oxidation rates and increased glucose oxidation rates in the hearts of mice lacking PPARalpha are associated with higher concentrations of malonyl-CoA and reduced expression of malonyl-CoA decarboxylase. J. Biol. Chem. 277,4098-4103[Abstract/Free Full Text]
  35. Lankford, E. B., Epstein, N. D., Fananapazir, L., Sweeney, H. L. (1995) Abnormal contractile properties of muscle fibers expressing beta-myosin heavy chain gene mutations in patients with hypertrophic cardiomyopathy. J. Clin. Invest. 95,1409-1414[Medline]
  36. Lowey, S. (2002) Functional consequences of mutations in the myosin heavy chain at sites implicated in familial hypertrophic cardiomyopathy. Trends Cardiovasc. Med. 12,348-354[CrossRef][Medline]
  37. Szczesna, D., Ghosh, D., Li, Q., Gomes, A. V., Guzman, G., Arana, C., Zhi, G., Stull, J. T., Potter, J. D. (2001) Familial hypertrophic cardiomyopathy mutations in the regulatory light chains of myosin affect their structure, Ca2+ binding, and phosphorylation. J. Biol. Chem. 276,7086-7092[Abstract/Free Full Text]
  38. Holroyde, M. J., Potter, J. D., Solaro, R. J. (1979) The calcium binding properties of phosphorylated and unphosphorylated cardiac and skeletal myosins. J. Biol. Chem. 254,6478-6482[Abstract/Free Full Text]
  39. Da Silva, A. C., Kendrick-Jones, J., Reinach, F. C. (1992) Hybrid myosin light chains containing a calcium-specific site from troponin C. Eur. J. Biochem. 204,85-91[Medline]
  40. Reinach, F. C., Nagai, K., Kendrick-Jones, J. (1986) Site-directed mutagenesis of the regulatory light-chain Ca2+/Mg2+ binding site and its role in hybrid myosins. Nature 322,80-83[CrossRef][Medline]
  41. Kendrick-Jones, J., Rasera da Silva, A. C., Reinach, F. C., Messer, N., Rowe, T., McLaughlin, P. (1991) Recombinant DNA approaches to study the role of the regulatory light chains (RLC) using scallop myosin as a test system. J. Cell Sci. Suppl. 14,55-58[Medline]
  42. Da Silva, A. C., Kendrick Jones, J., Reinach, F. C. (1992) Hybrid myosin light chains containing a calcium-specific site from troponin C. Eur. J. Biochem. 204,85-91[Medline]
  43. Szczesna, D., Sobieszek, A., Kakol, I. (1987) Binding of phosphorylated and dephosphorylated heavy meromyosin to F-actin. FEBS Lett. 210,177-180[CrossRef][Medline]
  44. Diffee, G. M., Greaser, M. L., Reinach, F. C., Moss, R. L. (1995) Effects of a non-divalent cation binding mutant of myosin regulatory light chain on tension generation in skinned skeletal muscle fibers. Biophys. J. 68,1443-1452[Medline]



This article has been cited by other articles:


Home page
 Cardiovasc ResHome page
T. P. Abraham, M. Jones, K. Kazmierczak, H.-Y. Liang, A. C. Pinheiro, C. S. Wagg, G. D. Lopaschuk, and D. Szczesna-Cordary
Diastolic dysfunction in familial hypertrophic cardiomyopathy transgenic model mice
Cardiovasc Res, April 1, 2009; 82(1): 84 - 92.
[Abstract] [Full Text] [PDF]

Home page
 FASEB J.Home page
W. G. L. Kerrick, K. Kazmierczak, Y. Xu, Y. Wang, and D. Szczesna-Cordary
Malignant familial hypertrophic cardiomyopathy D166V mutation in the ventricular myosin regulatory light chain causes profound effects in skinned and intact papillary muscle fibers from transgenic mice
FASEB J, March 1, 2009; 23(3): 855 - 865.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
fj.07-8630comv1
21/14/3974    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Szczesna-Cordary, D.
Right arrow Articles by Lopaschuk, G. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Szczesna-Cordary, D.
Right arrow Articles by Lopaschuk, G. D.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS