Three-dimensional structure of vertebrate cardiac
muscle myosin filaments
Maria E. Zoghbi*, John L. Woodhead*, Richard L. Moss†, and Roger Craig*‡
*Department of Cell Biology, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, MA 01655; and†Department of Physiology,
University of Wisconsin School of Medicine and Public Health, Madison, WI 53706
Edited by Hugh E. Huxley, Brandeis University, Waltham, MA, and approved December 13, 2007 (received for review September 19, 2007)
Contraction of the heart results from interaction of the myosin and
actin filaments. Cardiac myosin filaments consist of the molecular
motor myosin II, the sarcomeric template protein, titin, and the
cardiac modulatory protein, myosin binding protein C (MyBP-C).
Inherited hypertrophic cardiomyopathy (HCM) is a disease caused
mainly by mutations in these proteins. The structure of cardiac
myosin filaments and the alterations caused by HCM mutations are
unknown. We have used electron microscopy and image analysis
to determine the three-dimensional structure of myosin filaments
from wild-type mouse cardiac muscle and from a MyBP-C knockout
model for HCM. Three-dimensional reconstruction of the wild-type
filament reveals the conformation of the myosin heads and the
organization of titin and MyBP-C at 4 nm resolution. Myosin heads
appear to interact with each other intramolecularly, as in off-state
smooth muscle myosin [Wendt T, Taylor D, Trybus KM, Taylor K
(2001) Proc Natl Acad Sci USA 98:4361–4366], suggesting that all
relaxed muscle myosin IIs may adopt this conformation. Titin
domains run in an elongated strand along the filament surface,
where they appear to interact with part of MyBP-C and with the
myosin backbone. In the knockout filament, some of the myosin
head interactions are disrupted, suggesting that MyBP-C is impor-
tant for normal relaxation of the filament. These observations
provide key insights into the role of the myosin filament in cardiac
contraction, assembly, and disease. The techniques we have de-
veloped should be useful in studying the structural basis of other
myosin-related HCM diseases.
electron microscopy ? MyBP-C ? thick filament ? three-dimensional
ments that fill cardiac muscle cells. The thick filaments are
polymers of myosin II together with associated proteins, includ-
ing titin and myosin binding protein C (MyBP-C) (1). Mutations
in myosin and MyBP-C are the most common cause of inherited
hypertrophic cardiomyopathy (HCM), a relatively common and
often fatal disease characterized by left ventricular hypertrophy,
myocyte disarray, and interstitial fibrosis (2, 3). The molecular
mechanisms by which HCM mutations lead to the pathogenesis
of this disease are unclear.
Structural knowledge of vertebrate thick filaments in the
relaxed state has come from x-ray diffraction and electron
microscopy (EM) of striated muscle (4–9). Myosin molecules
assemble into bipolar filaments, with their ?-helical coiled-coil
tails in the filament backbone and their paired heads on the
surface, in three near-helical strands having a repeat of ?43 nm
and an axial rise between levels of heads of ?14.3 nm (1, 4, 9).
Antiparallel overlap between myosin tails at the center of the
filament creates a ‘‘bare zone,’’ free of myosin heads, where
filament polarity reverses. Three-dimensional EM studies have
revealed the near-helical distribution of the myosin heads in
skeletal muscle in the relaxed state. However, conformational
details have been limited by the low resolution of the recon-
structions, and neither MyBP-C nor titin have been resolved
ontraction of the heart depends on interactions between the
thick (myosin-containing) and thin (actin-containing) fila-
(10–12). In addition, three-dimensional reconstructions of car-
diac filaments have not been reported.
MyBP-C (13) is located at seven to eight axial sites, 43 nm
apart (the same as the myosin repeat), in the middle third of each
half of the filament (14–16), where it plays a role in modulating
Ig-like (Ig) and fibronectin-like (Fn) domains, linearly arranged,
but its detailed organization on the filament is unknown. Some
observations suggest that the C-terminal of MyBP-C wraps
around the filament backbone (15, 17), whereas others suggest
that its C-terminal runs parallel to the filament axis with its
N-terminal extending out toward the surrounding thin filaments
(15, 18). Titin is a giant (3 MDa) protein extending from the
center of the filament to the Z line (14). In the thick filament
region, it consists of a long string of Ig and Fn modules arranged
in repeating patterns that bind to myosin and MyBP-C and
appear to correlate with the ?43-nm periodicity of these com-
ponents (19). These and other observations suggest that titin is
a developmental template for sarcomere assembly (14, 19, 20),
but its organization on the filament (whether it is located in the
filament core or on its surface, and whether helically or linearly
detailed molecular organization of myosin, titin and MyBP-C
that underlies normal functioning of the thick filament, there is
little information on how mutations in these proteins cause
Here, we present a three-dimensional reconstruction of a
cardiac thick filament, revealing the structure of the myosin
heads in the relaxed state and the organization of titin and
MyBP-C. Comparison with a reconstruction of filaments from a
MyBP-C knockout mouse model for HCM (21) provides addi-
tional evidence for the location of MyBP-C and reveals the
impact of its loss on thick filament structure. The results provide
important insights into normal thick filament function and
assembly, and into how alterations in thick filament structure
may affect cardiac muscle function.
Three-Dimensional Reconstruction Reveals the Near-Helical Array of
Myosin Heads. Thick filaments isolated from wild-type mouse
ventricle appeared intact by negative staining electron micros-
copy (Fig. 1 A and B). Fourier transforms indicated good
preservation of the ordered array of myosin heads (Fig. 1C), with
reflections at the expected positions for a perturbed (8, 9, 22, 23)
Author contributions: M.E.Z. and R.C. designed research; M.E.Z. performed research;
M.E.Z., J.L.W., and R.L.M. contributed new reagents/analytic tools; M.E.Z., J.L.W., and R.C.
analyzed data; and M.E.Z., J.L.W., and R.C. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The data reported in this paper have been deposited in the EBI Macro-
molecular Structure Database, www.ebi.ac.uk (accession no. EMD-1465).
‡To whom correspondence should be addressed. E-mail: firstname.lastname@example.org.
This article contains supporting information online at www.pnas.org/cgi/content/full/
© 2008 by The National Academy of Sciences of the USA
February 19, 2008 ?
vol. 105 ?
no. 7 www.pnas.org?cgi?doi?10.1073?pnas.0708912105
helical structure with a 42.9-nm repeat. Three-dimensional
reconstruction of the MyBP-C region of the filament (approx-
imately the middle third of each half) was carried out by single
particle methods (10, 24) [see Materials and Methods and sup-
porting information (SI) Methods]. The reconstruction, based on
2564 segments, had a resolution of 3.2–4.0 nm according to the
of two better than previous vertebrate thick filament reconstruc-
tions, all from skeletal muscle [6.5 nm (10), according to the
same FSC criterion, and ?7 nm (12)]. At this resolution, key new
structural features become visible. The main surface features are
crown has 3-fold rotational symmetry (Fig. 1 D and E and SI
Movie 1). These features must represent myosin heads, the main
globular components in the thick filament. The head array
exhibits a perturbed helical structure, consistent with previous
3 are close to their correct axial (14.3 nm) and azimuthal helical
positions. However, crown 2 is axially closer to crown 3 (13.2 nm)
than to crown 1 (15.4 nm), and is at almost the same azimuthal
position as crown 1 (instead of being rotated by 40°, as expected
for a perfect helix; SI Fig. 8). The axial perturbations are similar
to those seen in a reconstruction of fish skeletal filaments (10)
and in a 2D analysis of rabbit cardiac filaments (23).
Atomic Fitting Reveals Intramolecular Interaction Between Myosin
Heads. The enhanced resolution of the reconstruction makes
possible a relatively precise fitting of myosin head atomic
models. The striking visual resemblance, in two dimensions,
between the motifs at crowns 1 and 2 and the atomic model of
regulated myosin in the enzymatically inactive (‘‘off’’) state (24,
25) (SI Fig. 9) suggested that this was an appropriate model for
3D fitting. In this model, the heads are in the ADP.Pi state, as
are the myosin filaments under the conditions of our experi-
ments. We found that this model gave an excellent and unam-
biguous 3D fit into the globular features in crowns 1 and 2 (Fig.
2 and SI Movie 2). The myosin heads appear to interact
asymmetrically, with their motor domains pointing toward the
filament bare zone. This conclusion is supported by the obser-
vation of similar asymmetric head-head interactions in isolated,
negatively stained mouse cardiac myosin molecules under relax-
ing conditions (H. Jung and R.C., unpublished data), and by the
observation of a folded conformation (suggestive of head–head
interaction) in rotary shadowed cardiac myosin molecules (26).
Attempts to dock the two-headed model (as well as individual
ADP.Pi-like S1 molecules) in other orientations were unsuccess-
ful. Thus cardiac myosin, which is thought to have no intrinsic
Electron micrograph of negatively stained filaments. (Scale bar, 500 nm.) (B)
Portion of isolated filament, with bare zone at top. (Scale bar, 43 nm.) (C)
Fourier transform of B showing layer lines at orders (numbered) of 42.9-nm
repeat. (D and E) Surface views of thick filament reconstruction (filtered to
4.0-nm resolution), showing a single 42.9-nm repeat (scale bar), oriented so
levels (crowns 1, 2, and 3) in each repeat.
Isolation and 3D reconstruction of mouse cardiac thick filaments. (A)
(24) to head motif in crowns 1 and 2 of reconstruction. The 3D envelope has
been made translucent to aid visualization of enclosed atomic model. Small
regions of atomic model outside the envelope appear bright, whereas the
majority, enclosed within it, appears dull (see also SI Movie 2). The heads are
labeled ‘‘blocked’’ and ‘‘free’’ according to terminology proposed to describe
the actin-binding capability of each head for regulated myosin (24, 25).
‘‘Blocked’’ head: motor domain, green; essential light chain, orange; regula-
tory light chain, yellow. ‘‘Free’’ head: motor domain, cyan; essential light
chain, pink; regulatory light chain, beige. Bare zone direction toward top.
Fitting of atomic model of myosin heads in off-state conformation
Zoghbi et al.
February 19, 2008 ?
vol. 105 ?
no. 7 ?
regulation and therefore to lack an off-state (27), adopts a
similar conformation to the off-state of regulated myosin (see
discussion in ref. 24).
cannot be determined unambiguously from the reconstruction.
Attempts to fit the two-headed off-state model, or atomic
models of individual myosin heads (e.g., ref. 28), were unsuc-
cessful because the reconstructed volume is greater than that
required for one head, but too small for two. The smaller
apparent volume of crown 3 heads suggests that they may have
greater mobility than those in crowns 1 and 2.
The Reconstruction Reveals the Organization of Titin and MyBP-C on
the Filament Surface. The reconstruction also has features that
cannot be attributed to myosin heads. These are therefore likely
to be myosin-binding proteins, whose molecular organization on
the filament has not been previously visualized. In the space
between azimuthally adjacent pairs of heads are small transverse
bands that are especially evident in crowns 1 and 2 (blue coloring
in Fig. 3B; SI Movies 1 and 3). Eleven such densities (arrows in
Fig. 3A), ?4 nm apart axially, are seen in each 42.9-nm repeat.
These appear to form a longitudinal strand running approxi-
mately parallel to the filament axis, at a radius of 7–8 nm (blue
arrows, Fig. 3 C–E), on the surface of the backbone. A similar
periodicity has also been observed in a 2D analysis of cardiac
myosin filaments (22). The arrangement of these 4 nm ‘‘beads’’
is consistent with the linear organization of the globular, 4 nm
previously been inferred only from sequence data. Six molecules
of titin are expected for each half of the thick filament, possibly
arranged in pairs (29). However, our data do not allow us to
distinguish any particular arrangement of the two putative titin
molecules within a pair. The near azimuthal alignment of crowns
1 and 2 allows titin to follow a linear course in this region (Fig.
3B). The marked curvature around crown 3 might contribute to
the different properties of the myosin heads in this crown.
than the putative titin density (orange arrow in Fig. 3D). This
density corresponds to three additional features, 4 nm apart,
observed in the longitudinal surface view of the filament (orange
coloring in Fig. 3B), to the left of the putative titin strand. The
axial position of this additional density coincides with the
location of strong density seen in striated muscle A-bands, which
is known to represent MyBP-C (SI Methods and SI Fig. 10). The
three 4-nm-diameter domains are consistent with the linear
organization of Fn and Ig domains in MyBP-C. They may
represent domains of MyBP-C known to interact with titin
and/or the myosin tail (15).
Three-Dimensional Reconstruction of MyBP-C Knockout Filament. We
have also carried out a 3D reconstruction of cardiac thick
filaments from a MyBP-C knockout mouse model for HCM (21)
(SI Methods). Although the overall appearance of wild-type and
knockout filaments appears to be similar (SI Fig. 11; ref. 30), the
diameter of the knockout reconstruction and the mass of the
heads appear slightly smaller, and the resolution is lower (?7
nm, compared with 4 nm for the wild-type), suggesting that the
array of heads is less stable in filaments lacking MyBP-C. In
addition, there is a clear structural change in crown 1 of the
knockout filaments (Fig. 4 and SI Movie 4), such that the heads
In contrast, crowns 2 and 3 appear relatively unaffected by the
absence of MyBP-C. The restriction of the main structural
change to crown 1 adds support to our conclusion that MyBP-C
is in this crown. It also suggests that MyBP-C may be important
heads in this crown.
Intramolecular Interaction Between Myosin Heads in the Cardiac Thick
Filament. The conformation of the myosin heads and putative
organization of titin and MyBP-C that we have observed provide
of reconstruction (rear side removed for clarity). Arrows point to 11 densities
spaced 4 nm apart within the 42.9-nm repeat (protein white). (Scale bar, 43
nm.) (B) Surface view of same region at higher contour level, showing the
three crowns of myosin heads (green). Surface features corresponding to the
strand of densities in A are visible (blue, representing titin), together with
three additional densities in crown 1 (orange, thought to represent MyBP-C).
(C–E) Projections of transverse sections of the central regions of crowns 2, 1,
and 3 (where the strand in A runs roughly parallel to the filament axis),
showing the radial positions of the densities in A (blue arrows) and of the
three additional beads of density in crown 1 in B (orange arrow). Note that,
because the filaments have 3-fold rotational symmetry, each of the features
marked in A–E is repeated at azimuthal angles of 120° and 240° with respect
to those shown.
thick filaments. (A) Wild type. (B) Knockout. The reconstructions are based on
a similar number of segments in each case. Both have been filtered to 7-nm
resolution (the resolution of the knockout filaments) to enable a direct
comparison. Circle shows altered structure of myosin heads in crown 1 of
knockout. (Scale bar, 43 nm.)
www.pnas.org?cgi?doi?10.1073?pnas.0708912105 Zoghbi et al.
important new insights into the structure and function of ver-
tebrate cardiac muscle thick filaments. Previous structural anal-
whereas 3D reconstructions, carried out on vertebrate skeletal
thick filaments [from frog (12) and fish (10) muscle] had
considerably lower resolution (?7 nm). The 3- to 4-nm resolu-
tion of the current work makes possible a rather precise fitting
of myosin head atomic models. A surprising finding is the
excellent fit of the interacting-head conformation of regulated
myosin heads in the off-state (24, 25) to two of the three crowns
of the 42.9-nm repeat of the filament. Head–head interaction
was previously thought to be the mechanism by which ATPase
and actin-binding activity were switched off in intrinsically
regulated myosins (e.g., vertebrate smooth, invertebrate striated
muscle myosin) (24, 25). One head (‘‘blocked;’’ Fig. 2) would be
prevented from binding to actin and the other head (‘‘free’’)
would be unable to hydrolyze ATP (24, 25). Myosin from
vertebrate-striated muscles is thought to lack an intrinsic regu-
latory switch and is therefore thought to be unable to switch off
biochemically (these muscles are regulated primarily through the
troponin–tropomyosin system on the thin filaments) (27). The
observation of the interacting-head motif in cardiac filaments
would therefore suggest that head-head interaction per se does
not switch activity off. It may instead be a resting position or
‘‘relaxed state’’ conformation common to both regulated and
unregulated myosin filaments. In unregulated myosins, the head-
head interaction may be weak and thus may not inhibit myosin
function (although it might restrict head access to actin filaments
in the relaxed state; ref. 31). Additional interactions present in
regulated myosins (e.g., involving the regulatory light chains, or
interactions with heads in other crowns or with S2; ref. 24) may
by stabilizing the interacting-head structure. Alternatively, ver-
tebrate thick filaments might possess a regulatory mechanism
that has not yet been detected biochemically.
Whereas the heads in crowns 1 and 2 have a ‘‘relaxed
conformation,’’ those in crown 3 appear to be more mobile. In
the sarcomere, where actin filaments are close to the thick
filaments, greater freedom of crown 3 heads may increase their
probability of interaction with actin. Two populations of heads
may allow fine-tuning of thick filament activity. Activity is also
modulated by phosphorylation of the regulatory light chains,
which increases head mobility and enhances contraction in both
vertebrate skeletal and cardiac muscle (32, 33), and by MyBP-C
phosphorylation does not merely modulate activity, but fully
activates switched off molecules. In both systems, phosphoryla-
tion probably functions by breaking the head–head (and other)
interactions that maintain the relaxed conformation (25).
Titin Is Linearly Organized on the Thick Filament Surface. The im-
proved resolution of our reconstruction has enabled the visual-
ization of nonmyosin thick filament proteins whose organization
was previously unknown. The linear arrangement of eleven,
4-nm-diameter globular features in each 42.9-nm repeat directly
suggests that we are visualizing titin (14, 35). The 11 domains are
likely to correspond to the 43-nm-long superrepeat of seven Fn
and four Ig domains of this size in titin in the MyBP-C region of
the filament, previously identified on the basis of sequence data
and thought to dictate the location of myosin and MyBP-C (14).
The titin strands lie on the surface of the filament backbone
(consistent with titin epitope availability in antibody labeling
studies; ref. 36), oriented approximately parallel to the filament
axis, where they can form longitudinal interactions with myosin
tails and MyBP-C. Our observations appear to rule out models
in which titin follows the helical path of the myosin heads or is
buried in the filament core. They suggest instead that titin forms
a simple linear scaffold for filament formation, and that myosin
molecules assemble on the interior of this scaffold during
sarcomere development and turnover. Interestingly, there ap-
pears to be no simple matching of the axial positions of myosin
heads (spaced 15.2, 13.4, and 14.3 nm apart) with titin domains
(?3.9 nm apart), as might have been expected from the template
function of titin. Thus, during thick filament assembly, a differ-
ent set of titin–myosin interfaces must be used at each of the
Organization of MyBP-C on the Filament Surface. The molecular
organization of MyBP-C in the thick filament has remained a
mystery since its discovery 35 years ago (13, 15), despite its
emerging importance as a modulator of cardiac contraction and
as a prime target of mutations causing HCM. Our results suggest
that at least three of its 11 Fn and Ig domains run along the
filament surface parallel to the axis (18), interacting with titin
and probably with myosin tails in crown 1. These interactions,
presumably involving the C-terminal region of MyBP-C (known
to interact with myosin tails and titin; ref. 15), may anchor
MyBP-C to the thick filament. We see no evidence that its other
domains form a collar around the backbone, as has been
suggested (15), although such an arrangement should be visible
if present. They may instead extend from the filament surface
(18) at crown 1 where they could bind to subfragment 2 and/or
actin, modulating myosin–actin interaction and force generation
(15). We observe MyBP-C stripes only in sarcomeres with a very
well ordered lattice (SI Fig. 10), suggesting that any such
extended regions might be disordered in isolated filaments,
explaining their absence from our reconstruction.
Implications from MyBP-C Knockout Filament. MyBP-C is important
for normal functioning of the heart (15) and appears to play a
‘‘cardioprotective’’ role in ischemic injury (37). Mouse models are
being used to understand the functioning of MyBP-C (38). Knock-
out mice demonstrate that MyBP-C is not essential for survival,
although its absence causes cardiac hypertrophy (21). The similar
overall structure of MyBP-C knockout and wild type filaments (SI
Fig. 11 A and B, compare Fig. 1 A and B) may explain the viability
of these mice and the relatively good prognosis for most HCM
patients carrying MyBP-C mutations (38). The lower resolution of
the MyBP-C knockout reconstruction suggests that MyBP-C helps
to stabilize the relaxed array of myosin heads in normal filaments
(see also ref. 30). The decreased ability of crown 1 heads to adopt
the interacting-head conformation in the absence of MyBP-C
suggests that the knockout filaments may not relax as fully as wild
type. Failure of the filaments to relax properly in the absence of
MyBP-C may offer a molecular explanation of the compromised
relaxation of the heart in MyBP-C knockout mice (21, 39) and may
also be related to the abnormal diastolic function in humans with
Our reconstructions show that cardiac thick filaments are
complex structures, where not all myosin heads behave in a
proteins. This diversity suggests that cardiac thick filament
function may be finely modulated. The techniques we have
developed here open the way to further structural studies on
HCM mouse models carrying different mutations in MyBP-C or
other thick filament proteins. Such studies should reveal whether
alteration in the relaxed conformation of the thick filaments is
a common molecular mechanism underlying HCM.
Materials and Methods
mice (8–10 weeks old, strain 129 SVE, Taconic Farms, NY). Mice were injected
i.p. with heparin (1,000 units/kg) and 15 min later anesthetized with sodium
pentobarbital (60 units/kg). Once anesthetized, mice were killed by cervical
dislocation. These procedures were approved by the University of Massachu-
setts Medical School Institutional Animal Care and Use Committee.
Zoghbi et al.
February 19, 2008 ?
vol. 105 ?
no. 7 ?
Immediately after euthanasia, the heart was harvested and placed in
calcium-free Tyrode (140 mM NaCl, 5.4 mM KCl, 1.5 mM MgCl2, 10 mM Hepes,
0.33 mM Na2HPO4, 10 mM glucose, pH 7.4) plus 20 mM 2,3-butanedione
monoxime (BDM, to enhance actin–myosin dissociation). The aorta was cath-
eterized for retrograde perfusion of the coronary arteries with the following
solutions: first, calcium-free Tyrode plus 20 mM BDM for 5 min; second,
relaxing solution (100 mM NaCl, 2 mM EGTA, 5 mM MgCl2, 1 mM DTT, 10 mM
8 min; third, relaxing solution for 5 min. After this final wash, the ventricles
were cut and homogenized in relaxing solution for 10 sec at position 5 using
a Polytron homogenizer (Brinkmann Instruments, Westbury, NY). The whole
procedure was carried out at room temperature.
During homogenization in relaxing solution, only a small number of fila-
of a method used for insect flight muscle (40). The homogenate was spun at
10,000 g for 2 min to obtain a pellet of myofibrils, which were resuspended
and incubated with 70 ?g calpain-1 (Calbiochem) in 48 ?l 20 mM imidazole-
HCl, 5 mM 2-mercaptoethanol, 1 mM EDTA, 1 mM EGTA, 30% glycerol, pH 6.8
plus 10 ?M free Ca2?for 1 h at room temperature. Enzymatic treatment was
stopped by addition of relaxing solution, containing 100 ?M blebbistatin
(Toronto Research Chemicals), and the suspension was agitated to help re-
lease thick and thin filaments (blebbistatin was used to aid release of thick
filaments and to stabilize the relaxed conformation of the heads (F. Zhao and
R.C., unpublished data). Thin filaments were removed by fragmentation with
gelsolin (41). The thick filament suspension was kept on ice and used within
24 h. For the rationale for these procedures, see SI Methods.
Electron Microscopy. The ordering of myosin heads is rapidly lost in mamma-
by negative staining at near-physiological temperature. This avoided difficul-
provides a very good representation of protein distribution in most macro-
molecular structures (43), including thick filaments (1). A drop of filament
of carbon supported by a thicker holey carbon film. The grid was rinsed
EGTA, 5 mM imidazole, 1 mM sodium azide, 1 mM MgATP, pH 7.0) and five
drops of 2% uranyl acetate. Staining was carried out at room temperature
with solutions prewarmed to 37°C. Grids were observed in a Philips CM120
electron microscope (FEI, Hillsboro, OR) at 80 KV under low dose conditions.
Images of filaments on thin carbon over holes were acquired at 42,000?
magnification, using a 2Kx2K CCD camera (F224HD, TVIPS GmbH, Gauting,
Germany) at a resolution of 0.57 nm/pixel.
Image Analysis. Long, straight thick filaments, with a clearly visible bare zone,
were chosen for analysis. Filaments were oriented vertically, with the bare
zone at the top, and the region between the 3rd and 10th 42.9-nm repeats
from the bare zone (where MyBP-C is present) was computationally cut
single-particle analysis, we used only those regions where the Fourier trans-
forms showed the layer lines expected for a well ordered vertebrate thick
Selected filament regions were converted to SPIDER format (EM2EM; Im-
was carried out by using SPIDER (see below). UCSF Chimera (beta version 1,
build 2199) was used for visualization, analysis, and atomic fitting of 3D
volumes (44) (see SI Methods).
Three-Dimensional Reconstruction. Single particle analysis rather than Fourier-
Bessel based helical methods were used for 3D reconstruction so that the
known perturbations in the helical order of vertebrate myosin filaments
would be preserved and not averaged out (11) (see SI Methods for details).
Electron micrograph images of filaments are 2D projections perpendicular to
the filament axis. Because filaments are expected to lie on the EM grid with
different rotations about their long axes, micrographs of different filaments
should represent different rotational views. If the relative rotational angle of
each view is known, the 3D structure of the filaments can be determined by
back projection using those angles. Relative rotations of different filament
segments were determined by matching filament images against 2D projec-
tions of 3D models rotated around their long axis at known angles (see SI
that it most closely matched, and the views were back-projected to generate
and the process was iterated until there were no further changes in the
reconstruction (10, 24).
Possible model bias in the reconstruction was ruled out by our finding that
the final reconstruction was the same with three different, independent
starting models (see SI Methods; and SI Fig. 6). For true helical structures,
helical refinement can improve the signal:noise ratio and resolution of the
reconstruction (24, 45). We avoided helical refinement to preserve the helical
perturbations present in vertebrate filaments (9, 10, 12, 23). However, C3
symmetry was imposed to take advantage of their threefold rotational sym-
for filament isolation. This work was supported by an American Heart Asso-
ciation Postdoctoral Fellowship (to M.E.Z.) and National Institutes of Health
Grants AR34711 (to R.C.) and HL82900 (to R.L.M.). Electron microscopy was
carried out in the Core Electron Microscopy Facility of the University of
Massachusetts Medical School, supported in part by Diabetes Endocrinology
Research Center Grant DK32520. Molecular graphics images were produced
by using the UCSF Chimera package from the Resource for Biocomputing,
Visualization, and Informatics at the University of California, San Francisco
(supported by National Institutes of Health Grant P41 RR-01081).
1. Craig R, Woodhead JL (2006) Curr Opin Struct Biol 16:204–212.
2. Marian AJ (2005) Curr Cardiol Rev 1:53–63.
3. Fatkin D, Graham RM (2002) Physiol Rev 82:945–980.
4. Huxley HE (1963) J Mol Biol 7:281–308.
5. Kensler RW, Stewart M (1983) J Cell Biol 96:1797–1802.
6. Kensler RW, Stewart M (1989) J Cell Sci 94:391–401.
7. Kensler RW, Stewart M (1993) J Cell Sci 105:841–848.
8. Kensler RW (2002) Biophys J 82:1497–1508.
9. Huxley HE, Brown W (1967) J Mol Biol 30:383–434.
10. Al-Khayat HA, Morris EP, Kensler RW, Squire JM (2006) J Struct Biol 155:202–217.
11. Eakins F, Al-Khayat HA, Kensler RW, Morris EP, Squire JM (2002) J Struct Biol 137:154–
12. Stewart M, Kensler RW (1986) J Mol Biol 192:831–851.
13. Offer G, Moos C, Starr R (1973) J Mol Biol 74:653–676.
14. Gregorio CC, Granzier H, Sorimachi H, Labeit S (1999) Curr Opin Cell Biol 11:18–25.
15. Flashman E, Redwood C, Moolman-Smook J, Watkins H (2004) Circ Res 94:1279–1289.
16. Bennett P, Craig R, Starr R, Offer G (1986) J Muscle Res Cell Motil 7:550–567.
17. Moolman-Smook J, Flashman E, de Lange W, Li Z, Corfield V, Redwood C, Watkins H
(2002) Circ Res 91:704–711.
18. Squire JM, Luther PK, Knupp C (2003) J Mol Biol 331:713–724.
19. Labeit S, Gautel M, Lakey A, Trinick J (1992) EMBO J 11:1711–1716.
20. Tskhovrebova L, Trinick J (2003) Nat Rev Mol Cell Biol 4:679–689.
21. Harris SP, Bartley CR, Hacker TA, McDonald KS, Douglas PS, Greaser ML, Powers PA,
Moss RL (2002) Circ Res 90:594–601.
22. Kensler RW (2005) J Struct Biol 149:313–324.
23. Kensler RW (2005) J Struct Biol 149:303–312.
24. Woodhead JL, Zhao FQ, Craig R, Egelman EH, Alamo L, Padron R (2005) Nature
25. Wendt T, Taylor D, Trybus KM, Taylor K (2001) Proc Natl Acad Sci USA 98:4361–4366.
26. Takahashi T, Fukukawa C, Naraoka C, Katoh T, Yazawa M (1999) J Biochem (Tokyo)
27. Lehman W, Szent-Gyorgyi AG (1975) J Gen Physiol 66:1–30.
28. Dominguez R, Freyzon Y, Trybus KM, Cohen C (1998) Cell 94:559–571.
29. Liversage AD, Holmes D, Knight PJ, Tskhovrebova L, Trinick J (2001) J Mol Biol
30. Kensler RW, Harris SP (2007) Biophys J 94, in press.
31. Zoghbi ME, Woodhead JL, Craig R, Padron R (2004) J Mol Biol 342:1223–1236.
32. Sweeney HL, Bowman BF, Stull JT (1993) Am J Physiol 264:C1085–C1095.
33. Levine RJ, Kensler RW, Yang Z, Stull JT, Sweeney HL (1996) Biophys J 71:898–907.
34. Oakley CE, Chamoun J, Brown LJ, Hambly BD (2007) Int J Biochem Cell Biol 39:2161–
35. Trinick J, Knight P, Whiting A (1984) J Mol Biol 180:331–356.
36. Whiting A, Wardale J, Trinick J (1989) J Mol Biol 205:263–268.
Seidman JG, Robbins J (2006) Proc Natl Acad Sci USA 103:16918–16923.
38. Tardiff JC (2005) Heart Fail Rev 10:237–248.
39. Pohlmann L, Kroger I, Vignier N, Schlossarek S, Kramer E, Coirault C, Sultan KR,
El-Armouche A, Winegrad S, Eschenhagen T, et al. (2007) Circ Res 101:928–938.
40. Kulke M, Neagoe C, Kolmerer B, Minajeva A, Hinssen H, Bullard B, Linke WA (2001)
J Cell Biol 154:1045–1057.
41. Hidalgo C, Padron R, Horowitz R, Zhao FQ, Craig R (2001) Biophys J 81:2817–2826.
42. Xu S, Offer G, Gu J, White HD, Yu LC (2003) Biochemistry 42:390–401.
43. Ohi M, Li Y, Cheng Y, Walz T (2004) Biol Proced Online 6:23–34.
44. Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE
(2004) J Comput Chem 25:1605–1612.
45. Egelman EH (2000) Ultramicroscopy 85:225–234.
www.pnas.org?cgi?doi?10.1073?pnas.0708912105Zoghbi et al.