5. L. P. Jackson et al., Cell 141, 1220–1229 (2010).
6. L. M. Traub, Nat. Rev. Mol. Cell Biol. 10, 583–596 (2009).
7. E. M. Schmid, H. T. McMahon, Nature
448, 883–888 (2007).
8. M. J. Taylor, D. Perrais, C. J. Merrifield, PLOS Biol. 9, e1000604 (2011).
9. E. ter Haar, S. C. Harrison, T. Kirchhausen, Proc. Natl. Acad.
Sci. U.S.A. 97, 1096–1100 (2000).
10. M. A. Edeling et al., Dev. Cell 10, 329–342 (2006).
11. Materials and methods are available as supplementary
materials on Science Online.
12. S. Zaremba, J. H. Keen, J. Cell Biol. 97, 1339–1347 (1983).
13. R. Lindner, E. Ungewickell, J. Biol. Chem. 267, 16567–16573
14. S. Höning et al., Mol. Cell 18, 519–531 (2005).
15. J. Hirst et al., Mol. Biol. Cell 24, 2558–2569 (2013).
16. L. M. Traub, J. A. Ostrom, S. Kornfeld, J. Cell Biol. 123,
17. X. Ren, G. G. Farías, B. J. Canagarajah, J. S. Bonifacino,
J. H. Hurley, Cell 152, 755–767 (2013).
18. M. Beck et al., Mol. Syst. Biol. 7, 549 (2011).
19. D. Loerke et al., PLOS Biol. 7, e57 (2009).
20. G. H. Borner et al., J. Cell Biol. 197, 141–160 (2012).
21. A. Motley, N. A. Bright, M. N. Seaman, M. S. Robinson, J. Cell
Biol. 162, 909–918 (2003).
22. A. P. Liu, F. Aguet, G. Danuser, S. L. Schmid, J. Cell Biol. 191,
23. F. M. Brodsky, Annu. Rev. Cell Dev. Biol. 28, 309–336 (2012).
24. F. Aguet, C. N. Antonescu, M. Mettlen, S. L. Schmid,
G. Danuser, Dev. Cell 26, 279–291 (2013).
25. R. A. Laskowski, M. B. Swindells, J. Chem. Inf. Model. 51, 2778–
We would like to thank the I02, I03, and I04-1 beamline staff at the
Diamond Lightsource; A. McCoy and P. Evans for crystallographic
advice; C. Oubridge for advice and assistance with SeMet mapping
of the hinge residues; N. Bright, C. Savva, and S. Miller for advice
and assistance with EM; C. Smith for reagents; and H. Böning and
H. Ungewickell for expert technical assistance. D.J.O. and B. T.K.
are supported by a Wellcome Trust Principal Research fellowship
(090909/Z/09/Z). S.H. is supported by a grant of the German
Science Foundation (SFB 635, TP A3). S.C.G. is supported by a
Sir Henry Dale fellowship from the Wellcome Trust and the Royal
Society (098406/Z/12/Z). CIMR is supported by a Wellcome
Trust Strategic Award (079895). Coordinates have been deposited
in the Protein Data Bank with PDB ID 4uqi.
Materials and Methods
Figs. S1 to S11
Tables S1 and S2
Movies S1 and S2
15 April 2014; accepted 24 June 2014
Mechanism of actin filament pointed-end
capping by tropomodulin
Jampani Nageswara Rao, Yadaiah Madasu, Roberto Dominguez*
Proteins that cap the ends of the actin filament are essential regulators of cytoskeleton
dynamics. Whereas several proteins cap the rapidly growing barbed end, tropomodulin
(Tmod) is the only protein known to cap the slowly growing pointed end. The lack of
structural information severely limits our understanding of Tmod’s capping mechanism.
We describe crystal structures of actin complexes with the unstructured amino-terminal
and the leucine-rich repeat carboxy-terminal domains of Tmod. The structures and
biochemical analysis of structure-inspired mutants showed that one Tmod molecule
interacts with three actin subunits at the pointed end, while also contacting two
tropomyosin molecules on each side of the filament. We found that Tmod achieves
high-affinity binding through several discrete low-affinity interactions, which suggests
a mechanism for controlled subunit exchange at the pointed end.
The proteins that cap the ends of the actin filament play important roles in actin-driven processes such as cell migration and or- ganelle trafficking by controlling the ad- dition and dissociation of actin subunits
at filament ends. Several proteins cap the barbed
end of the filament, including capping protein
(CP) and some gelsolin-family members (1, 2).
In contrast, tropomodulin (Tmod) is the only
protein known to cap the pointed end of tropomyosin (TM)–coated actin filaments (3). Four
Tmod isoforms work in conjunction with one
of several TM isoforms to stabilize actin structures characterized by a uniform distribution
of the lengths of actin filaments. These structures include the sarcomere of cardiac and skeletal muscle cells and the spectrin-based membrane
skeleton (3, 4).
The mechanism by which Tmod caps the
pointed end is poorly understood. Quantification
in skeletal muscle and erythrocytes led to the pro-
posal that two Tmod molecules cap the pointed
end (5, 6). In vitro, however, one Tmod molecule
is sufficient to block pointed-end elongation of
TM-coated filaments (7), consistent with the dis-
tinctive domain architecture of Tmod, which har-
bors two actin- and two TM-binding sites. Thus,
the N-terminal ~160–amino acid region is mostly
unstructured in isolation (8), but contains three
predicted helical segments that bind TM, actin,
and TM in that order (9, 10). This region displays
TM-dependent capping activity (9). Most of the
C-terminal region (human Tmod1 residues 161
to 359) consists of a leucine-rich repeat (LRR)
domain (11). This region displays limited cap-
ping activity on its own (9). Although Tmod binds
with nanomolar affinity to the pointed end (12),
and even greater affinity in the presence of TM
(7), it does not form an absolute cap. Instead,
Tmod functions as a “leaky” cap, determining
the length of the actin filaments while allow-
ing for the controlled addition or dissociation
of actin subunits at the pointed end (13). In the
absence of high-resolution structures of the
pointed end, rationalization of the existing data
is difficult, and several models exist, featuring
either one (10, 11) or two (5, 6) Tmod molecules
at the pointed end.
Actin polymerization prevents crystallization
of capping complexes. We thus attempted crystal-
lization of the N- and C-terminal actin-binding
sites (ABS1 and ABS2) of Tmod in complex with
monomeric actin. However, both sites bound
with weak affinity to monomeric actin (see be-
low), and polymerization persisted during crys-
tallization. A solution was found by fusing ABS1
and ABS2 C-terminally to gelsolin segment 1 (GS1)
via a nine–amino acid flexible linker [crystalliza-
tion strategies are described in (14)]. The Tmod
fragments extended beyond the actin-binding
sites defined previously (9, 15–17), with ABS1 and
ABS2 comprising human Tmod1 residues 50 to
101 and 160 to 349, respectively (Fig. 1A). No-
tably, ABS1 and ABS2 both bound actin:GS1 with
1:1 stoichiometry and with similar affinities [dis-
sociation constant (KD) = 7.5 and 10.5 mM for
ABS1 and ABS2, respectively] when not connected
by a linker (Fig. 1, B and C). Similar binding
affinities were obtained at two different tem-
peratures, 10°C and 20°C, and with adenosine
5´-triphosphate (ATP)– or adenosine 5´-diphosphate
(ADP)–actin (fig. S1).
The complexes of ATP-actin with GS1-ABS1
and GS1-ABS2 crystallized under slightly different conditions, and with different unit cell
parameters (14) (table S1). The structures were
determined to 1.8 and 2.3 Å resolution for ABS1
and ABS2, respectively (Fig. 1, D and E, and fig.
S2, A and B). Both structures were well defined
in the electron density maps (fig. S2, C and D).
The flexible linkers between GS1 and the Tmod
fragments, and residues 50 to 57 and 100 to 101
of ABS1 and 160 to 169 of ABS2, were not visualized. These residues likely do not interact
with actin, because the C termini of ABS1 and
ABS2 projected away from actin, and weak electron density that could not be modeled was also
observed projecting away from actin at their N
termini. A structure of ABS1 was also obtained
with ADP-actin at 2.15 Å resolution and showed
a conformation similar to that observed with
ATP-actin (14). Crystals of ABS2 could not be
obtained with ADP-actin.
The Tmod structures differed substantially
from previously characterized actin complexes
(18); whereas most actin-binding proteins bind
in the cleft between actin subdomains 1 and 3,
ABS1 and ABS2 bound at the pointed end of the
actin monomer, and on the side of subdomains
Department of Physiology, Perelman School of Medicine,
University of Pennsylvania, Philadelphia, PA 19104, USA.
*Corresponding author: E-mail: firstname.lastname@example.org