Effects of hypertrophic and dilated cardiomyopathy mutations on
power output by human
James A. Spudich1,*, Tural Aksel1, Sadie R. Bartholomew1, Suman Nag1, Masataka Kawana1,
Elizabeth Choe Yu1, Saswata S. Sarkar1, Jongmin Sung1, Ruth F. Sommese1, Shirley Sutton1, Carol Cho1, Arjun S. Adhikari1, Rebecca Taylor1, Chao Liu1, Darshan Trivedi1and Kathleen M. Ruppel1,2,*
Hypertrophic cardiomyopathy is the most frequently occurring inherited cardiovascular disease, with a prevalence of more than one in 500 individuals worldwide. Genetically acquired dilated cardiomyopathy is a related disease that is less prevalent. Both are caused by mutations in the genes encoding the fundamental force-generating protein machinery of the cardiac muscle sarcomere, including human β-cardiac myosin, the motor protein that powers ventricular contraction. Despite numerous studies, most performed with non-human or non-cardiac myosin, there is no clear consensus about the mechanism of action of these mutations on the function of humanβ-cardiac myosin. We are using a recombinantly expressed humanβ-cardiac myosin motor domain along with conventional and new methodologies to characterize the forces and velocities of the mutant myosins compared with wild type. Our studies are extending beyond myosin interactions with pure actin filaments to include the interaction of myosin with regulated actin filaments containing tropomyosin and troponin, the roles of regulatory light chain phosphorylation on the functions of the system, and the possible roles of myosin binding protein-C and titin, important regulatory components of both cardiac and skeletal muscles.
KEY WORDS: Cardiac myosin, Cardiomyopathy mutations, Force–velocity curves
Dozens of laboratories, working over the course of more than a century and using a wide variety of approaches, have contributed importantly to our understanding of muscle contraction (Rall, 2014). The muscle sarcomere may well be the most studied protein complex in cell biology, partially due to its extraordinarily high degree of structural organization.
I (J.A.S.) was asked to preface my talk at the 2015 Journal of Experimental Biology Symposium on Muscle: Molecules to Motion with a brief historical perspective of my laboratory’s specific contributions to research on actin and myosin, and how those contributions led to our current focus on understanding the molecular basis of hypertrophic and dilated cardiomyopathy. My laboratory, over the last several decades, has been devoted to developing bothin vivoandin vitrosystems to understand the molecular basis of energy transduction by the myosin family of molecular motors (Spudich, 2011, 2012). Our interests have included the roles of myosins in non-muscle cells and the molecular basis of non-muscle contraction. Essential
for our research was the development ofin vitromotility assays for purified actin and myosin in the 1980s (Kron and Spudich, 1986; Spudich et al., 1985). We used these assays to show that the globular head of myosin, known as subfragment-1 (S1), is the motor domain of the myosin molecule (Toyoshima et al., 1987), and, together with molecular genetic approaches, we provided early strong functional evidence that the light-chain binding region of the S1 acts as a swinging lever arm during chemo-mechanical coupling (e.g. Shih et al., 2000; Uyeda et al., 1996). Our study on changing the lever arm length and even removing the light-chain binding region altogether was an early work that showed that the heavy chain, truncated after the converter domain (Fig. 1, myosin head structure on the right), is the ‘motor domain’of the molecule, and the lever arm amplifies allosteric movements in the motor domain to obtain a larger stroke size–the distance myosin moves an actin filament per ATP hydrolyzed (Uyeda et al., 1996). Using the physics of laser trapping, we simplified the motility assay to the single molecule level, which allowed the measurement of the stroke size (∼10 nm) as well as the intrinsic force produced by the motor (a few piconewtons) (Finer et al., 1994). With these tools in hand, and after studying a variety of myosin motor types over several decades, my laboratory is now fully focused on one of the most clinically important members of the myosin family of molecular motors, humanβ-cardiac myosin.
Human cardiomyopathy mutations are a leading cause of cardiac death
The humanβ-cardiac myosin motor is a major site in the sarcomere for hundreds of missense mutations that result in a spectrum of heart diseases called cardiomyopathies. Genetically based hypertrophic cardiomyopathy (HCM), an autosomal dominant inherited disease (Geisterfer-Lowrance et al., 1990; Seidman and Seidman, 2000, 2001), is clinically characterized by thickening of the ventricular walls with a resultant decrease in ventricular chamber size that occurs without a predisposing cause. Systolic performance of the heart is preserved or even increased, but relaxation capacity is diminished. HCM affects more than one in 500 individuals (Harvey and Leinwand, 2011; Maron, 2011; Maron et al., 1995; Semsarian et al., 2015). Dilated cardiomyopathy (DCM), in contrast, is characterized by thinning of the heart walls, enlargement of the ventricular chambers, and weakness of the cardiac muscle, resulting in an inability to raise the cardiac output to meet the demands of even modest exercise (Hershberger et al., 2013).
There are reported to be more than 300 pathogenic mutations in
β-cardiac myosin (Buvoli et al., 2008; Colegrave and Peckham, 2014; Seidman and Seidman, 2001; Walsh et al., 2010). Most of these are HCM mutations in the globular head domain, myosin S1 (Colegrave and Peckham, 2014). The other major site in the sarcomere for HCM-causing mutations is a regulatory protein known as myosin binding protein-C (MyBP-C). MyBP-C is thought
Department of Biochemistry, Stanford University School of Medicine, Stanford, CA 94305, USA.2Department of Pediatrics (Cardiology), Stanford University School of Medicine, Stanford, CA 94305, USA.
*Authors for correspondence ( email@example.com; firstname.lastname@example.org)
to put a break on the contraction of the muscle, which can be regulated by its phosphorylation (Chow et al., 2014; Kensler et al., 2011; Seidman and Seidman, 2000; van Dijk et al., 2014). A smaller but significant set of mutations are found in the tropomyosin– troponin complex. Tropomyosin (Tm) and the three troponin subunits, troponin T (TnT), troponin I (TnI), and troponin C (TnC), form the Ca2+-regulatory complex of the sarcomere. The actin,
myosin, tropomyosin, and the three troponins constitute a six-component system that is considered to be the fundamental regulated contractile complex of the sarcomere (Fig. 2). MyBP-C, which is in the vicinity of the myosin heads in the sarcomere, should be added as a seventh component of the fundamental regulated complex. Furthermore, the giant protein titin is also in the vicinity of the myosin heads in the sarcomere and has been shown to carry DCM mutations (Gerull et al., 2002; Herman et al., 2012; LeWinter and Granzier, 2013, 2014).
Much more is known about the role of the Tm.Tn regulatory complex in cardiac contraction than that of the regulatory proteins MyBP-C and titin. There is good evidence that domains of MyBP-C near the N terminus bind to the regulated actin filament while its C terminus binds to the myosin thick filament backbone (Fig. 2), thus playing a regulatory role in muscle contraction (Chow et al., 2014; Kensler et al., 2011; van Dijk et al., 2014). Studies also suggest that
MyBP-C binds to myosin S2, the coiled-coil domain just C-terminal to the globular S1 head, as well as to the regulatory light chain (RLC) of the myosin S1 (Harris et al., 2004; Kampourakis et al., 2014; Pfuhl and Gautel, 2012; Ratti et al., 2011). Titin has been thought of as an element involved in passive tension in the sarcomere (Granzier and Irving, 1995), but as discussed below, MyBP-C and titin may both contribute importantly to the regulation of ensemble force production by the myosin.
The force–velocity curve is fundamental to muscle contraction
As described above, clinically HCM mutations are associated with hypercontractility, whereas DCM mutations are associated with hypocontractility. This has led to the general hypothesis that HCM mutations are hypercontractile and DCM mutations are hypocontractile at the fundamental molecular level. Many important studies have contributed to our current understanding of the effects of some HCM and DCM mutations on various functions of the purified actomyosin complex, usually its ATPase activity or its velocity in thein vitromotility assay (for review, see Moore et al., 2012; Sivaramakrishnan et al., 2009). Only a few studies have attempted to explore the effects of the mutations on the power output generated by a reconstituted actin–myosin complex (Aksel et al., 2015; Debold et al., 2007; Greenberg et al., 2010; Sommese et al., 2013b). Ultimately, the molecular basis of changes in power output is what one needs to understand.
Power (P) output comes directly from the force–velocity (F–V) curve of muscle contraction, asP=F×v(Fig. 3). The force axis on this curve is related to the load on the muscle; the force the muscle produces must overcome the load. Ventricular function is influenced by the two types of loads that are applied to cardiac muscle. One is preload, the load applied by the volume of blood filling the left ventricle that results in stretching of the myofibrils prior to the initiation of ejection. The other is afterload, the load due to pressure in the systemic circulation during contraction when the aortic valve is open. The afterload is the primary load that the heart needs to overcome to eject blood out of the left ventricle.
A somewhat oversimplified but useful view of velocity (v) and ensemble force (Fensemble) breaks these down into five parameters,f,
d,ts,tc, andNt(Fig. 3). The velocity at any load is a reflection ofd/ts,
the distance moved per strongly-bound state time of the motor to actin. Fensemble is the intrinsic force f of each motor domain
multiplied by the total number of myosin heads in a force-producing
0 2 4 6 8 10 12 14
Lever arm length (nm) measured from putative fulcrum point
0 1 2 3 4
Sliding velocity (
Fig. 1. Sliding velocity as a function of lever arm length.TheDictyosteium discoideum myosin II gene was genetically engineered to produce four forms of S1 with different lever arm lengths (shown on the right), expressed in D. discoideum, and purified as described previously (Uyeda et al., 1996). The normal D. discoideummyosin S1 has two light chains. Constructs containing zero light chains and one light chain were expressed, as well as one containing three light chains. The latter was created by inserting an additional essential light chain (magenta) binding domain into the myosin II heavy chain. The fulcrum was inferred from the intercept of the straight line with thex-axis.vo,
unloaded sliding velocity;d, stroke size;ts,
strongly bound state time. Data are from Uyeda et al. (1996).
List of symbols and abbreviations
DCM dilated cardiomyopathy ELC essential light chain
f intrinsic force
Fensemble ensemble force
HCM hypertrophic cardiomyopathy MyBP-C myosin binding protein-C
Nt total number of functionally available heads
RLC regulatory light chain
sS1 short subfragment-1 tc total cycle time
TnC troponin C
TnI troponin I
TnT troponin T
ts strongly bound state time
WT wild type
state at any moment. The latter is the total number of heads functionally available (Nt) times the duty ratio. The duty ratio for
those heads available to interact with the actin ists/tc, wheretsis the
strongly bound state time of myosin on the actin at a particular load and tc is the total cycle time of the actin-activated myosin
chemomechanical cycle. The weakly bound states of the motor are the ATP- and ADP-Pi-bound states, while the strongly bound states are the ADP-bound states (Fig. 4). Thus the ensemble force in the sarcomere isFensemble=f(ts/tc)Nt.
We have been examining these individual parameters for several myosin HCM and DCM mutations studied in a recombinant form of humanβ-cardiac myosin (short S1 or sS1) that consists of residues 1–808 (the motor domain elongated by the essential light chain [ELC] binding domain) co-expressed with the human cardiac ventricular ELC (MYL3) (Fig. 1, second myosin head image).
Changes in intrinsic force, cycle time and strongly bound state time are generally small for HCM and DCM mutations
Although several-fold changes in intrinsic force, cycle time or strongly bound state time have been reported in studies of HCM and DCM using non-human myosins compared with wild type (WT), we have found only small (10–50%) changes in any of these parameters for the HCM mutant forms we have studied using the human β-cardiac sS1 (R403Q, R453C, R719W, R723G, and R663H). In general, these relatively small changes are not surprising, as most patients with HCM do not begin to exhibit symptoms until at least the third decade of life (and often much later).
The DCM mutants (I201T, A223T, R237W, S532P, and F764L) showed somewhat larger changes compared with WT than the HCM mutants. M531R, a left ventricular non-compaction mutant, showed the largest change, thetcof which was decreased 1.7-fold compared
with WT (Aksel et al., 2015).
In general, we are finding that the DCM mutations all behave similarly; for example, they all increasetc. In contrast, the HCM
mutations are variable, some appearing more hypocontractile with regard to these parameters and others slightly hypercontractile. These variabilities led us to develop a metric that relates toFensemble
more directly, using a loadedin vitromotility assay.
An ensemble force metric derived from a loadedin vitro motility assay
One way to measure actin gliding velocities at different loads is using a loaded in vitro motility assay in which increasing concentrations of an actin binding protein is utilized as the load-bearing molecule (Bing et al., 2000; Greenberg and Moore, 2010;
Percent velocity Percent power
Fig. 3. Schematic plot of a force–velocity (F–V) curve for heart muscle. The power curve is generated by multiplying the force times the velocity at each point along the F–V curve. The grey zone is the region of the power curve thought to dominate systolic contraction.
Tn complex Tm Actin
Fig. 2. Schematic drawing to scale of essential components of the sarcomere.Tropomyosin (Tm) and the three subunits of troponin (Tn complex) decorate the actin filament. The domains of myosin are the S1 head (only one of two are shown for clarity), the coiled-coil S2, and the light meromyosin that forms the thick filament (bottom grey platform). The N-terminal domains of myosin binding protein-C (MyBP-C) bind to the thin actin filament and the C-terminal domains bind to the thick myosin filament. Titin (not shown) traverses in between the thin and thick filaments, closely associated with the thick filament backbone. Interactions among titin, MyBP-C, S1, and S2 are possible and perhaps regulatory, but have not been fully explored.
5. ATP hydrolysis
3. ADP release 4. ATP binding
2. Power stroke Pi release
1. Actin binding T
Fig. 4. Schematic figure of the key steps of the actin-activated myosin chemomechanical cycle.The total cycle involves five key steps. The pre-stroke states of S1 are shown on the right and the post-pre-stroke states on the left. Step 1: The pre-stroke S1 with bound ADP (D) and phosphate (Pi) undergoes a
weak to strong transition in binding to actin. This is the rate-limiting step in the cycle and determines the total cycle time (tc), which is 1/kcat, wherekcatis the
turnover number, or the number of substrate molecules each enzyme site converts to product per unit time. Step 2: While tightly bound to actin, the lever arm swings about its fulcrum point (black circle) to the right to its post-stroke position (black arrow), moving the actin filament to the left (bold blue arrow) with respect to the myosin thick filament. Step 3: ADP release frees the active site for binding of ATP (T). This step is related to the velocity of the actin filament sliding as the filament cannot move any faster than the myosin heads can let go. Step 4: ATP binding weakens the interaction of the S1 to actin. Although the ATP-bound heads still maintain a weak affinity for the actin, this state is typically drawn as dissociated from the actin filament. Step 5: ATP hydrolysis results in cocking of the head into the pre-stroke state (Shih et al., 2000).
Haeberle and Hemric, 1995; Warshaw et al., 1990). This assay has been used to compare power output and maximal force generation between myosin variants (Greenberg et al., 2010; Haeberle, 1994; Malmqvist et al., 2004) and between different solution conditions (Greenberg and Moore, 2010; Janson et al., 1992). The load-bearing molecule often used isα-actinin.
Theα-actinin assay does not work well, however, with the small motor domain sS1, presumably because of the small size of this domain compared with that of two-headed myosin constructs (Aksel et al., 2015). To obtain information about the relative ensemble forces of mutant and WT human β-cardiac sS1, we developed an utrophin-based loadedin vitromotility assay (Aksel et al., 2015) (Fig. 5A). The actin-binding protein utrophin is used to put a load on the actin filaments that the myosin must overcome. To achieve proper control of myosin and load molecule ratios, a truncated utrophin construct of similar size to sS1 is engineered to anchor on a glass surface via the same attachment used for sS1. Both the velocity of the filaments and the percentage of mobile filaments decrease as utrophin concentration is increased. These two parameters can be combined into what we call the percent time mobile (Fig. 5B). This parameter is a proxy for the ensemble force generated by the motor molecule (Aksel et al., 2015). Importantly, to facilitate these experiments, we also developed a rapid, objective filament-tracking software called FAST (Aksel et al., 2015). (For details of the FAST classes and routines, see the FAST package available at http://spudlab.stanford.edu/fast-for-automatic-motility-measurements/. FAST is currently available for both Linux
[Ubuntu 14.04] and Mac operating systems. Academic license for FAST is available for free on http://spudlab.stanford.edu/fast-for-automatic-motility-measurements/.)
We have used the utrophin loaded motility assay and FAST to characterize humanα- andβ-cardiac sS1 (Aksel et al., 2015). As seen in Fig. 5B, the amount of utrophin needed to overcome the motility of human α-cardiac sS1 is considerably less than that needed to overcome the humanβ-cardiac sS1, consistent withα -cardiac sS1 being a weaker motor ( producing lessFensemble) than β-cardiac sS1 (Malmqvist et al., 2004). We have also applied this approach to mutants that show opposite mechanical and enzymatic phenotypes with respect to each other, the hypercontractile left ventricular non-compaction mutant M531R and the hypocontractile DCM mutant S532P (Aksel et al., 2015). By this test, M531R is a stronger motor than WT, while S532P is a weaker motor. We also showed that omecamtiv mecarbil, a previously discovered cardiac specific myosin activator (Malik et al., 2011), increases
β-cardiac sS1Fensemblegeneration as judged by this loadedin vitro
motility assay (Aksel et al., 2015). We are now applying this approach to more than 15 cardiomyopathy mutant forms of human
HCM and DCM mutations in the tropomyosin–troponin complex are Ca2+-sensitizing and Ca2+-desensitizing,
The tropomyosin–troponin complex is the Ca2+ sensor in the
muscle and is composed of tropomyosin and three troponin subunits: (1) TnC, which binds Ca2+; (2) TnI, which binds actin
and tropomyosin and is the inhibitory subunit; and (3) TnT, which binds tropomyosin and communicates the Ca2+signal from
TnC to tropomyosin (Brown and Cohen, 2005; Kobayashi and Solaro, 2005). In the absence of Ca2+, tropomyosin is in a
position along the actin filament that blocks the binding of myosin. Upon Ca2+ binding to TnC, the inhibitory region of TnI
dissociates from tropomyosin, allowing the tropomyosin to move azimuthally on the actin filament, revealing the myosin binding sites on actin. Mutations in these proteins have been studied using various methods, including: animal models, skinned fibers into which mutant proteins have been exchanged, and reconstitutedin vitro molecular studies using endogenous and/or recombinant proteins such as ATPase, fluorescence, and motility assays (e.g. Miller et al., 2001; Mirza et al., 2005).
Extensive studies by many investigators have yielded important advances in understanding the effects of HCM- and DCM-causing mutations on a reconstituted six-component system consisting of actin, myosin, tropomyosin, and the three troponin subunits (for reviews, see Hitchcock-DeGregori, 2008; Kobayashi and Solaro, 2005; Moss et al., 2004; Tardiff, 2011; Willott et al., 2010). Most biochemical thin filament studies have been performed using non-human cardiac myosin (e.g. Szczesna et al., 2000; Venkatraman et al., 2005, 2003) or skeletal myosin (e.g. Kobayashi et al., 2013; Lin et al., 1996; Mirza et al., 2005; Tobacman et al., 1999). More recent studies have used human
β-cardiac myosin, where some specific differences in biochemical parameters were found compared with earlier non-human myosin studies (Bloemink et al., 2014; Deacon et al., 2012; Gupte et al., 2015; Sommese et al., 2013b). Regardless of the source of the mammalian myosin, however, the general paradigm holds true that HCM mutations in tropomyosin or troponin are sensitizers to Ca2+ (e.g. Fig. 6 shows HCM
mutations for three TnI mutations), while DCM mutations are desensitizers (Willott et al., 2010).
Human β-cardiac sS1
Utrophin concentration (nmol l–1)
ime mobile (%)
0 0.2 0.4 0.6 0.8
Fig. 5. Cartoon of the loadedin vitromotility assay and data obtained for humanα- andβ-cardiac sS1.(A) Myosin sS1 (red) and utrophin (green) are anchored to a microscope cover slip on the surface via the same attachment system involving binding of a C-terminal eight-residue peptide that specifically binds to SNAP-PDZ18 (light blue), which coats the surface. The utrophin puts a load on the actin filament and slows its gliding velocity. (B) Loadedin vitro motility percent time mobile data forα- (red) andβ- (blue) human cardiac sS1. Solid lines show the best stop-model fit (Aksel et al., 2015) to the percent time mobile data. Dashed lines mark values determined from the fit that reflect the relative force generation by the two motors; a lowerx-intercept means weaker
force. Data are from Aksel et al., (2015).
The myosin mesa and a possible role for MyBP-C and titin in regulating the number of functionally available myosin heads in the sarcomere
Besidesβ-cardiac myosin, the other major site in the sarcomere for HCM mutations is the regulatory protein MyBP-C (Buvoli et al., 2008; Seidman and Seidman, 2011; Walsh et al., 2010). MyBP-C is thought to put a break on the contraction of the muscle, which can be regulated by its phosphorylation (Chow et al., 2014; Kensler et al., 2011; Seidman and Seidman, 2000; van Dijk et al., 2014). There is good evidence that domains of MyBP-C near the N terminus bind to the regulated actin filament, whereas its C terminus binds to the myosin thick filament backbone (Fig. 2), playing a regulatory role in muscle contraction (Chow et al., 2014; Kensler et al., 2011; van Dijk et al., 2014). MyBP-C is in the A-band region of the sarcomere in close proximity to the myosin heads (Fig. 2). Another regulator, the giant molecule titin (Granzier and Irving, 1995; LeWinter and Granzier, 2013, 2014), extends through the A-band region, running
from the Z-disc, to which actin filaments are bound, to the M-line, the center of the sarcomere. Thus, each half sarcomere has titin molecules running its entire length, where the titin plays an important role in passive tension of the sarcomere (Granzier and Irving, 1995) as well as other roles (LeWinter and Granzier, 2014). A recent report made the suggestion that MyBP-C and/or other proteins in the vicinity of the myosin heads may play a role in sequestering some of the heads and taking them out of the functionally available pool, reducingNtand thusFensemble(Spudich,
2015). The concept of MyBP-C removing myosin heads from the functioning pool of heads in the sarcomere has been suggested previously, with evidence for binding of MyBP-C to S2 and the RLC (for reviews, see Kampourakis et al., 2014; Moss et al., 2015). But there may be a site on the motor domain of S1, which has previously been overlooked, that participates in such sequestration of myosin heads. This site has been called the myosin mesa, which is highly conserved among the cardiac myosins, from mouse α -cardiac to humanβ-cardiac myosin (Spudich, 2015). The mesa has many more arginine residues on its surface than lysine residues, the opposite of usual protein surfaces (Warwicker et al., 2014). Arginines are known to play a role in transient interactions between different proteins (Jones et al., 2000). Thus, the cluster of eight arginine residues on the mesa, R169, R204, R249, R403, R442, R453, R652, and R663, could be involved in transient interactions between the mesa and another protein or protein domain (e.g. MyBP-C, titin, myosin S2) (Fig. 7, blue-labeled amino acids). Strikingly, all eight of these arginines are sites of HCM mutations (Alfares et al., 2015). A number of other non-arginine cardiomyopathy mutations are also on the mesa in the vicinity of these arginines (Fig. 7). Besides a potential MyBP-C interaction with the myosin mesa, titin could be another player in such an interaction. Indeed, Granzier and colleagues reported that certain titin domains in the A-band of the sarcomere have conserved surface patterns and show weak interaction with myosin S1 (Muhle-Goll et al., 2001).
An intriguing hypothesis is that the myosin mesa HCM mutations reduce the affinity of S1’s putative binding partner(s) (e.g. MyBP-C, titin, myosin S2), releasing those heads to now be involved in the contractile process. This would result in hypercontractility of the
R453C R204H R249Q R442C R652G R663H
R453C R204H R249Q R442C R403Q
Fig. 7. Locations of 17 hypertrophic cardiomyopathy mutations in the myosin catalytic domain.(A) Cartoon showing two groups of HCM mutations. Four mutations are in the converter domain (dark green), and 13 are on or very near the myosin mesa (bright green). Sixteen of the HCM mutations were chosen because they have been documented to be the cause of HCM in families carrying these mutations (Alfares et al., 2015). The 17th is M531R, which, while only documented in one family, is a left ventricular non-compaction mutant myosin that appears to be hypercontractile in our studies. (B) The top view of the mesa showing 13 mutations on or near the mesa surface. The residues labeled in blue denote the positively charged amino acids. (C) The same view as in B except the surface charge distribution is shown. I263T, N444S, and M531R are slightly below the surface in acidic pockets, while the remainder of the mutations are on or very near the surface and most are arginine residues, seven of which form a particularly large domain of positive charge on the right half of the mesa as viewed.
8.0 7.5 7.0 6.5 6.0 5.5
Pase activity (%)
Fig. 6. Effects of TnI mutations on Ca2+regulation of actin-activated myosin S1 ATPase.Percent of the maximal wild-type (WT) ATPase activity (100%) is plotted as a function of the negative logarithm of the Ca2+ concentration. WT (black) is compared with R21C (blue), S166F (green) and
ΔK177 (red). Maximum WT activity was set to 100% and all other activities were normalized to this. Mean activities±standard error are plotted. Myosin S1 was prepared by proteolytic cleavage of bovine cardiac myosin.
muscle, which is characteristic of HCM clinically. Because the myosin motor domain is highly allosteric and there may be other binding sites for the putative interacting partner, this molecular mechanism for HCM hypercontractility may extend to other HCM mutations as well, and this mechanism could be a unifying hypothesis as to why HCM mutations are hypercontractile clinically.
Conclusion and future perspectives
Our laboratory is focused on changes in biochemical and biomechanical parameters in HCM and DCM mutant forms of humanβ-cardiac myosin. Our studies use both the two-component human system of actin and myosin (Aksel et al., 2015; Sommese et al., 2013b) and the six-component reconstituted human system: actin, myosin, Tm, TnC, TnI, and TnT (Gupte et al., 2015; Sommese et al., 2013a). From the considerations described here and elsewhere (Spudich, 2015), it is possible that many of the HCM mutations in the motor domain of myosin alter the effects of a putative MyBP-C, titin or myosin S2 interaction with the mesa surface. Thus, reconstitution of the eight-component system is likely to be necessary to fully understand the effects of the HCM mutations. This may equally apply to the DCM mutations.
Similarly, the effects of small molecule activators and inhibitors on the human cardiac ventricular system (e.g. Malik et al., 2011) will likely have to be understood at the eight-component level in molecular studies to correlate effects seen in the reconstituted system with those observed in skinned fibers, cardiomyocytes, the whole organ, and animals.
We thank Allan Borrayo for technical assistance in maintaining cell cultures and virus preparations.
J.A.S. is a founder of and owns shares in Cytokinetics, Inc., and MyoKardia, Inc., biotech companies that are developing therapeutics that target the sarcomere.
J.A.S. and K.M.R. developed the general concepts and approaches of the research program. T.A. developed the loadedin vitromotility assay. J.S. built the laser trap for single molecule analysis. S.N., M.K., A.A., S.S.S., T.A., J.S., R.F.S., S.R.B., E.C.Y., S.S., R.T., C.C., C.L., D.T., and K.M.R. performed experiments and, together with J.A.S., analyzed the data. S.R.B. carried out the experiments shown in Fig. 6. J.A.S. prepared the manuscript, which S.N., M.K., A.A., S.S.S., T.A., J.S., R.F.S., S.R.B., E.C.Y., S.S., R.T., C.C., C.L., D.T., and K.M.R. edited prior to submission.
This work was supported by the National Institutes of Health [GM33289 and HL117138 to J.A.S.]; the Human Frontiers Science Program [to J.A.S.]; a National Institutes of Health Cellular, Biochemical, and Molecular Sciences Training Grant [to S.R.B. and C.L.]; a Stanford Interdisciplinary Graduate Fellowship [to R.F.S.]; a Stanford Dean’s Postdoctoral Fellowship [to S.N.]; the National Science Foundation Graduate Research Fellowship Program [to S.R.B.]; and the Lucile Packard Child Health Research Institute Postdoctoral Grant [to A.S.A.]. Deposited in PMC for release after 12 months.
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