CORRECTED 1 APRIL 2021; SEE FULL TEXTSCIENCE ADVANCES | RESEARCH ARTICLE
BIOCHEMISTRY

The structure of the actin filament uncapping
complex mediated by twinfilin
Dennis M. Mwangangi1,2, Edward Manser1,2, Robert C. Robinson1,3,4*
Uncapping of actin filaments is essential for driving polymerization and depolymerization dynamics from capping protein–associated filaments; however, the mechanisms of uncapping leading to rapid disassembly are unknown. Here, we elucidated the x-ray crystal structure of the actin/twinfilin/capping protein complex to address
the mechanisms of twinfilin uncapping of actin filaments. The twinfilin/capping protein complex binds to two
G-actin subunits in an orientation that resembles the actin filament barbed end. This suggests an unanticipatedmechanism by which twinfilin disrupts the stable capping of actin filaments by inducing a G-actin conformationin the two terminal actin subunits. Furthermore, twinfilin disorders critical actin-capping protein interactions,which will assist in the dissociation of capping protein, and may promote filament uncapping through a second
mechanism involving V-1 competition for an actin-binding surface on capping protein. The extensive interactions
with capping protein indicate that the evolutionary conserved role of twinfilin is to uncap actin filaments.

Numerous cellular processes, such as morphogenesis, migration,cytokinesis, endocytosis, and memory, rely on rapid reorganizationof actin cytoskeletal networks (1–3). Coordinated local assemblyand disassembly of actin filaments generate force and structure,
which is harnessed to drive these specific functions. A number ofactin-regulating proteins control the assembly, disassembly, andorganization of the filament networks (4, 5). Among the key evolutionarily conserved actin regulators are capping protein (CP) andthe actin depolymerization factor homology (ADF-H) domain familyof proteins, which include ADF/cofilins and twinfilin (6–10). Whileprimitive, functional ADF-H domain proteins are found in Asgard
archaea (7, 9), CP and twinfilin have only been found, and are ubiquitous, in eukaryotes (9, 10). Thus, CP and twinfilin likely arose
during eukaryogenesis and were present in the last eukaryotic common
ancestor (LECA). The architecture of twinfilin is unique, composedof two ADF-H domains connected by a short linker and followed bya conserved C-terminal tail (11). ADF-H proteins generally regulate
cytoskeletal reorganization by accelerating the disassembly of actin
filaments (6, 12, 13). Twinfilin-1 is ubiquitously expressed in almost
all tissue types in mammals, where it regulates actin dynamics
through mechanisms involving interactions with actin monomers,
actin filaments, and CP (11, 14). The biological outputs from twinfilin regulation of actin dynamics include cell motility and synaptic
endocytosis (15).
The reported in vitro roles of twinfilin in actin dynamics are numerous, diverse, and somewhat contradictory. Twinfilin binds andsequesters adenosine diphosphate (ADP)–actin monomers withhigh affinity, inhibiting nucleotide exchange and preventing assembly into filaments (11, 14, 16). Twinfilin also interacts directly withactin filament barbed ends, blocking filament elongation, suggestive
1
Institute of Molecular and Cell Biology, A*STAR (Agency for Science, Technology
and Research), Biopolis, Singapore 138673, Singapore. 2Department of Pharmacology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore
117597, Singapore. 3School of Biomolecular Science and Engineering (BSE), Vidyasirimedhi Institute of Science and Technology (VISTEC), Rayong 21210, Thailand.
4
Research Institute for Interdisciplinary Science (RIIS), Okayama University, Okayama
700-8530, Japan.
*Corresponding author. Email: br.okayama.u@gmail.com

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of a capping activity (17–19). Recent studies have also demonstrated
that twinfilin accelerates depolymerization of actin filament barbed
ends containing ADP-actin subunits (20, 21), and at low pH, twinfilin can sever filaments (22). In addition, twinfilin interacts strongly
with heterodimeric CP through interactions that include those involving the conserved twinfilin C-terminal tail (23). CP is a hetero­dimer that binds to the barbed ends of actin filaments to prevent
actin subunit exchange (24–26). Although twinfilin binds to CP
with high affinity, its exact biological role in promoting CP cappingor uncapping is debated (27, 28). X-ray structural studies of twinfilin have been limited to single ADF-H domains, which show highstructural conservation, and both domains bind actin monomers(19). CP’s interactions with the actin filaments or dynactin filaments are resolved to 23- and 3.4-Å resolution, respectively, via
cryo–electron microscopy (cryo-EM) (29, 30). However, the molecular mechanism by which twinfilin interacts with CP at the actin
filament barbed ends is unknown. Here, we address the role oftwinfilin in uncapping of actin filaments by elucidating the x-ray
structure of the twinfilin/CP/actin complex.
RESULTSThe crystal structure of the twinfilin/CP/actin complex
Previous biochemical data have shown that twinfilin’s interaction
with CP-capped actin filaments protects CP from displacement by
CARMIL, suggesting a stable interaction between twinfilin, CP, andbarbed-end actin subunits (27). We therefore used purified twinfilin-1(human), heterodimeric CP (mouse CapZ1/2, henceforth CP),and skeletal muscle actin (rabbit) to reconstitute the complex between twinfilin, CP, and actin monomers. The twinfilin, CP, andactin complex was highly stable in gel filtration chromatography,
and this complex was used to prepare protein crystals suitable forstructure determination by x-ray crystallography at 3.2-Å resolution (fig. S1).
The complex consists of two ADP-bound actin subunits, one
subunit each of the heterodimeric CP (CP1 and CP2) and one
full-length twinfilin-1 (Fig. 1, A and B). In the structure, CP adopts
its canonical mushroom-shaped architecture consisting of a cap
and stalk and interacts with the barbed-end faces of two G-actin
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subunits via the top surface of the mushroom cap (Fig. 1). The two
actin subunits are in structurally similar conformations and adopt a
typical G-actin fold consisting of four subdomains. The actin subunits do not show the subunit flattening and twisting associatedwith the G-to-F actin transition (31). The deoxyribonuclease I binding loops in the two actin subunits are disordered (fig. S2, A and B).
The relative orientation of the two actin subunits resembles that ofbarbed-end actin subunits from the F-actin cryo-EM structure (fig.
S2, C to E) (32). The actin subunits are arranged by the short pitch
helical filament relationship, similar to two actin subunits across afilament. They do not adopt the relative positioning of two longitudinally related subunits in a single strand. Twinfilin adopts an elongated architecture in which the two ADF-H domains each bind one
actin subunit, with ADF-H domain 2 (D2) binding to the terminal
actin subunit, relative to a filament barbed end. The linker connecting the ADF-H domains, which includes an -helix, extends acrossthe upper surface (cap) of the mushroom-shaped CP and also contacts
both actin subunits (Fig. 1, A and B). The twinfilin C-terminal tailextends from D2 and wraps around the stalk of the CP -subunit,
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which is located opposite to the actin-binding interface, below theCP cap (Fig. 1, C and D).
Binary interactions in the twinfilin/CP/actin complex
Each of the two twinfilin ADF-H domains binds to an actin subunitat analogous interfaces, providing basis for monomer sequestration(Fig. 2A). The individual ADF-H domains adopt similar architectures except for a difference in the conformation of the -sheets.
The -3 and -4 strands in D2 form a protrusive extension relative
to that in ADF-H domain 1 (D1) (fig. S3, A to C). The key structuralelements of the two ADF-H domains are highly conserved in mousetwinfilin and human twinfilin-2 (33). Despite twinfilin D2 having
10 times higher affinity for G-actin (16), its actin-binding interface
is markedly similar to that of D1. Thus, the precise selection of residues in the two binding interfaces is likely to explain the differencesin actin-binding affinity. Further, the -helix in the linker between
D1 and D2 loosely associates with the D2-bound actin subunit, andthis likely strengthens D2 interaction with G-actin (Fig. 2A and fig.
S4, A and C).
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Fig. 1. The twinfilin/CP/actin complex. (A) Front view of the pentameric complex in cartoon representation. CP consists of two subunits, -subunit (CP) and -subunit(CP). Twinfilin comprises two ADF-H domains (D1 and D2), a linker between D1 and D2 that includes a helix (residues 151 to 165), and a C-terminal tail (Tail; residues 316
to 342, the last eight amino acids are disordered). Actin subunit 1 is bound to twinfilin D1, and subunit 2 to twinfilin D2. The twinfilin secondary structure elements arecolored differentially, helices in red, strands in cyan, and loops and extended regions in orange, with the Tail in lime green. (B) Front view [same as in (A)], in which theactin subunits and twinfilin are represented as surfaces. Twinfilin is shown entirely in orange. (C and D) Back and side views, respectively.

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The CP actin-binding interface is located at the top surface of themushroom cap (Fig. 2B). In the absence of twinfilin, CP interacts
with the barbed-end protomers of a filament via C-terminal extensions to the - and -subunits, the - and -tentacles (24, 26, 29). In
the twinfilin-bound structure, the CP -tentacle is mostly disordered in the structure and, hence, has no direct contact with eitherof the actin subunits (fig. S4, D and E). The -tentacle is partiallyordered and forms an interaction with actin 1 (fig. S4F). Twinfilinbinds CP through its C-terminal tail. This tail wraps around thestalk of CP -subunit, a common binding site for filament uncapping proteins with CP interaction (CPI) motifs (Fig. 2C) (34). The
ordered portion of the tail of twinfilin (residues 315 to 342) includes
a basic stretch that interacts with the negatively charged groove between the CP -subunit stalk and the underside of the mushroom-­
shaped cap (fig. S5, A to C). Furthermore, the complex reveals additional
interactions between twinfilin and CP, beyond the C-terminal tail.
First, the -helix in the linker between D1 and D2 forms an interaction with CP that also involves the start of the -tentacle (Fig. 2C
and fig. S5, D to F). Second, twinfilin D1 forms a direct interface
with the CP -subunit (Fig. 2C and fig. S5G).
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Comparison of the conformations of twinfilin tail–boundand CARMIL CPI–bound CP
The twinfilin tail-binding site on CP is distant from the actin-­binding site, which is centered on the -tentacle (Fig. 3, A and B).
Comparison of the tail-binding site with the uncapping CPI motif
from CARMIL shows an overlapping interaction on the undersideof the CP -subunit (34); however, the N termini of the two peptides take divergent paths around the CP stalk (Fig. 3A). The
CARMIL CPI motif half encircles the CP stalk, with its N terminus
making contact with the CP -subunit stalk. By contrast, the N terminus of the twinfilin tail (residues 316 to 322) follows a straightpath and does not form contacts with the CP -subunit. This region(residues 316 to 322) is elongated yet ordered with clear electron
density (Fig. 2C and fig. S5K), despite not being stabilized by interactions, suggesting that it may be under tension to extend the polypeptide chain into an ordered conformation. The overlapping
interface provides a structural basis for the competition between thetwinfilin tail and CARMIL CPI for CP binding, and twinfilin’s
attenuation of CARMIL-mediated dissociation of CP from filamentbarbed ends (27).
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Fig. 2. Binary interactions in the twinfilin/CP/actin complex. (A) Front and back views of actin bound to twinfilin D1 or twinfilin D2. (B) Two views of the CP interaction
with actin, in which actin 1 and actin 2 are shown in similar orientations. (C) Three orientations of the CP/twinfilin interaction. Examples of the electron density of key
features are shown in fig. S5 (H to K).

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CP can adopt two known conformations, which are likely to bein dynamic equilibrium in solution. The uncapping CPI motifs of
CARMIL, CD2AP, and CKIP stabilize one conformation, actin-free
conformation of CP (34). Superimposition of the complex withCARMIL-bound CP and unbound CP shows key structural changes
that CP subunits undergo in adopting the twinfilin/CP/actin complex conformation (Fig. 3, C and D, and fig. S6). First, the CP -subunit
mushroom cap in this complex moves upward relative to the stalk,while the -tentacle repositions to adopt the actin-bound conformation. Second, the -subunit adjusts upward flattening themushroom cap and adopts a similar conformation to that stabilizedby V-1 or Arp1 (from the dynactin complex) (Fig. 3E and fig. S6)(30, 35). The binding site of V-1, a steric CP inhibitor, overlaps withthat of actin in this complex (fig. S6J) (35). Therefore, the CP structure in the twinfilin/CP/actin complex represents the mushroom
cap–bound conformational state, to either actin, Arp1, or V-1,
while the CPI-bound structure of CP represents a stabilized mushroom cap–unbound conformation. Binding of the CARMIL CPI
motif around the stem of CP locks CP in the mushroom cap–unboundconformation, which is less compatible with actin barbed-end interaction, leading to uncapping of the filament (34, 36). The inability of the twinfilin tail to induce a change in the CP conformation
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from mushroom cap–bound to unbound state suggests that, in thecomplex, actin binding dominates the CP conformation or thetwinfilin tail does not stabilize the mushroom cap–unbound conformation. To distinguish between these possibilities, we tested for
uncapping activity within the twinfilin tail in isolation. We performed pyrene-actin polymerization assays in which an increase inpyrene fluorescence reports on the efficiency of actin assembly fromCP precapped actin filament seeds. The CARMIL CPI peptide, apositive control, displayed potent uncapping activity leading to polymerization from the CP precapped filament seeds, indicated by the increasein fluorescence (fig. S7). By contrast, the maltose-binding protein(MBP)–tagged twinfilin tail (residues Gln321-Asp350) had no detectable uncapping effect, showing a similar polymerization profile to
the CP precapped filament seeds alone (fig. S7). Thus, in this assay, thetwinfilin tail alone does not uncap CP-capped filaments, suggestingthat it does not strongly stabilize the CP mushroom cap–unboundconformation.
Twinfilin influences CP interaction with actin barbed ends
The structure reveals additional interactions between twinfilin and
CP that might influence the CP-binding mode to the actin filamentbarbed ends (fig. S5, D to G). The dynactin filament (consisting of
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Fig. 3. Conformations of CP. (A) Comparison of the twinfilin tail (yellow) binding site with CARMIL (cyan) on CP. Both CP-binding peptides run in the same direction, andN terminus of the twinfilin tail is labeled N. (B) The actin-binding site on the CP -tentacle is distant from the twinfilin tail-binding site. (C and D) Structural superimpositionof the -subunits of CP reveals that the conformation of CP in the twinfilin/CP/actin complex is different to the CARMIL-bound and unbound conformations of CP.
(E) Superimposition reveals that the conformation of CP in the twinfilin/CP/actin complex is similar to the V-1–bound conformation of CP. - and -subunits of CP arecolored light green and dark green, respectively, for the CARMIL, unbound, and V-1 complexes. Black arrows indicate conformational changes in CP in adopting thetwinfilin/CP/actin complex structure.

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This unstable interaction state may be targeted by other regulatory
factors, such as the CP-sequestering protein V-1. We used thepyrene-actin polymerization assay to test whether V-1 can enhance
CP uncapping in the presence of twinfilin (28). Addition of twinfilin
into CP precapped actin filament seeds mixed with pyrene-labeledmonomers and profilin did not accelerate polymerization, indicating either a lack of uncapping or uncapping followed by recapping(Fig. 5). Addition of V-1 alone displayed partial polymerization
attributable to CP sequestration (Fig. 5). However, the presence ofboth twinfilin and V-1 accelerated polymerization to near the levelof uncapped actin filament seeds, indicating that the cooperative
activities of twinfilin and V-1 can induce dissociation of CP frombarbed ends and prevent recapping by CP (Fig. 5). This indicatesthat twinfilin is able to remove CP from filaments; however, theCP-sequestering protein V-1 is required to prevent recapping of filaments. We propose that the high concentration of profilin (2.8 M),
relative to twinfilin (1 M), used in this assay was sufficient to competitively remove actin from the CP/twinfilin complex and allow
recapping in the absence of V-1. The requirement of both the uncapping agent twinfilin and CP-sequestering protein V-1 to observerobust uncapping in this in vitro assay partially explains some of thedisparities in the reported activities of twinfilin. Uncapping has
been difficult to observe in many assays in which recapping has notbeen excluded.
DISCUSSION

This analysis of the actin/twinfilin/CP complex reveals the structural
basis by which twinfilin interacts with actin and with CP, and provides insight into molecular mechanisms for the regulation of CP in
actin filament barbed-end dynamics. The unanticipated geometry

Fig. 4. Comparison of the CP-binding mode to Arp1 in the dynactin complex (Arp1/CP) with the binding mode to actin in actin/twinfilin/CP complex. (A andB) Focus on the - and -tentacles, respectively, which are highlighted by black circles. The -tentacle is disordered and unbound in the actin/twinfilin/CP complex, andthe -tentacle is partially bound and ordered, relative to the dynactin complex. Enlargements of the tentacle regions show the superimpositions of the CP (- and -subunits
colored red and blue, respectively) from the dynactin complex on to actin (green and teal) and twinfilin (yellow) from the actin/twinfilin/CP complex. (A) Enlargement, thetwinfilin linker (yellow) binds to the -tentacle–binding site on actin 2 (green). (B) Enlargement, the N terminus of twinfilin D1 (yellow) occupies half of the -tentacle–
binding site on actin 1 (teal).
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Arp1 instead of actin) is structurally similar to the actin filamentand is capped at its barbed end by CP (30). Comparison of theCP-binding modes to actin in the twinfilin/CP/actin complex and
to Arp1 subunits reveals similar overall geometries (Fig. 4 and fig.
S8). However, there are considerable differences in the binding of
the CP - and -tentacles to their respective actin/Arp1 subunits.
These differences arise from the presence of twinfilin, because thebinding modes of the - and -tentacles are similar in the dynactin
complex and in the 23-Å cryo-EM structure of the CP/actin filament (29, 30). The -tentacle, which is bound to the terminal Arp1
subunit in the dynactin complex, is disordered in the twinfilin/CP/
actin complex (Fig. 4A and fig. S8A). Structural superimpositionindicates that the twinfilin linker obscures the -tentacle bindingsite on actin in the actin/twinfilin/CP complex (Fig. 4A). In the dynactin complex, the -tentacle is fully ordered and bound to Arp1.
By contrast, the -tentacle in the twinfilin/CP/actin complex only
forms a partial interaction with actin (Fig. 4B and fig. S8, B and C).
Superimposition reveals that the -tentacle–binding site on actin ispartially obstructed by twinfilin D1 (Fig. 4B). The binding of thetentacles to the actin protomers will also be influenced by the actin
protomer conformation. A twinfilin-induced shift from an F-actin
to G-actin conformation may aid dissociation of the tentacles.
In the absence of twinfilin, the CP -tentacle is a critical interaction with barbed end of a filament, while the -tentacle offers a second important actin-binding interface to stabilize capping (36). The
obstruction of the CP tentacles by twinfilin in the actin/twinfilin/CP
complex will destabilize the CP interaction, leading to weakened CP
affinity for actin filament barbed ends. Thus, the CP-binding modein the dynactin complex represents fully bound CP, conferring strong
and stable capping activity, while the CP-binding mode in theactin/twinfilin/CP complex represents a weak and unstable capping activity.

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in which twinfilin binds to two actin subunits in a pseudo-filamentbarbed-end orientation has major implications for its possible mechanisms of action. Previously proposed mechanisms for twinfilin’s
activities assumed that twinfilin contacts two longitudinally related
actin subunits at the barbed end of an actin filament (17, 18, 21, 27, 28).
However, the structure presented here unambiguously identifies thatthe two actin subunits are laterally related. Twinfilin binds acrossthe two strands of an actin filament, rather than down the side of asingle strand in the filament. We discuss the ramifications of thisbinding orientation for actin filament regulation.
The primary biological role of twinfilin has not been settled.
Mammalian twinfilin-1 was shown to preferentially localize to regions of the cell enriched for F-actin (14). Loss of twinfilin-1 in B16
cells leads to their inability to generate lamellipodia (27). In vitro
and cell-based experiments have led to multiple proposed activities.
Twinfilin’s actin-related functions have been variously described asfollows: an actin monomer-sequestering protein that forms a 1:1
complex with actin that prevents actin polymerization and inhibits
nucleotide exchange in actin (11, 14); a CP-interacting and phosphatidylinositol 4,5-bisphosphate (PIP2)–interacting protein (37);an actin monomer-shuttling protein between the pointed and barbed
ends of filaments (38); a protein that does not affect CP-capping
activity, nor does CP affect twinfilin’s monomer binding function(rather, twinfilin localizes actin monomers to sites of assembly
through interaction with CP) (23); a barbed-end actin filament CP(17); an actin filament-severing protein (22); a protein that speeds
up actin depolymerization at both ends of the actin filament in conjunction with cyclase-associated protein (CAP) (20); a protein thatenhances CP actin filament capping, which competes with CARMIL
for CP binding (27); and an actin filament CP-uncapping protein(28). The actin/twinfilin/CP complex provides the structural basis
to reassess these activities.
In the complex, twinfilin forms extensive interactions with CP
and with two actin protomers, which adopt a pseudo-actin filament
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Fig. 5. V-1 uncaps filament barbed ends in the presence of twinfilin. Pyrene-actin
polymerization assay showing uncapping of F-actin seeds by V-1 in the presence of
twinfilin. Unlabeled F-actin seeds (0.5 M) were capped with 100 nM CP and subsequently polymerized in either 2 M actin (10% pyrene)/2.8 M profilin (red) or 2 Mactin (10% pyrene)/2.8 M profilin supplemented with either 1 M twinfilin (yellow),
5 M V-1 (purple), or a mixture of 1 M twinfilin and 5 M V-1 (blue). As a control,
F-actin seeds were polymerized in the absence of CP (black). As shown in the profile, the presence of both twinfilin and V-1 induces accelerated polymerization of precapped F-actin seeds (blue) to almost the same level as the positive control (black).
By contrast, twinfilin alone (yellow) induces very low level of polymerization, similar
to CP alone (red), while V-1 alone (purple) only induces minimal polymerization.
AU, arbitrary units.

barbed-end orientation relative to each other. These multiple interactions indicate that twinfilin’s primary and evolutionary conservedrole, throughout eukaryotes, is to regulate CP capping at the barbed
end of actin filaments. In yeast, the CP:twinfilin ratio is estimated tobe 2.5:1 (37, 39), which indicates that the actin-related cellular
activities of twinfilin will be dominated by the CP:twinfilin complex, rather than twinfilin acting alone on filaments. Similarly, themolar abundance of mammalian CP:twinfilin-1 ratio was determined as 2:1 (27). Twinfilin has been shown to strongly interact
with actin filament barbed ends and with CP, dissociation constants(Kd) of 13 and 50 nM, respectively (17, 27). However, these affinities
are at least one order of magnitude weaker than the CP affinity for
actin filament barbed ends [Kd = 0.1 to 1.0 nM (40)]. The actin/
twinfilin/CP complex structure demonstrates that twinfilin disrupts
CP-actin interfaces, sterically competing with the CP - and -­
tentacles in binding to actin, indicating that twinfilin’s principalrole is to destabilize CP capping (Fig. 6A). We discuss the possible
mechanisms that lead to uncapping.
The ADP-bound actin protomer conformations in the actin/
twinfilin/CP complex are very similar to that in the cofilin-decorated
actin filament (fig. S2B) (41). Cofilin, which consists of a single
ADF-H domain, severs actin filaments toward the pointed end of
a section of filament decorated with cofilin, by inducing a G-­
actin–like conformation in the actin protomers, which are not stable as a filament (13, 41). We hypothesize that the orientation of
the two ADF-H twinfilin domains, in binding across the filament,
allows twinfilin to induce a G-actin–like conformation in the final
two barbed-end actin protomers. This conformation will destabilize
these two terminal actin subunits, leading to them being “severed”
from the end of the filament, by an analogous mechanism to cofilin
severing (13, 41). Thus, we propose that the two twinfilin ADF
domains induce severing at the boundary between ADF-H–­
bound and ADF-H–free portions of F-actin, dissociating CP,
twinfilin, and the two terminal actin subunits as a complex (Figs. 1A
and 6, A and B). Immediate reassociation of the actin/twinfilin/
CP complex with the filament would be unfavorable because theactin subunits in the complex are held in the ADP-bound G-actin
state, and nucleotide exchange is inhibited by the twinfilin ADF-H
domains (11).
We propose that the actin/twinfilin/CP complex will then undergo a process of recycling. Twinfilin strongly binds to ADP-boundactin monomers (Kd ~ 40 to 50 nM), and this affinity is unaffectedby the presence of CP (16, 23). For comparison, CAP, thymosin-4,
profilin, and cofilin are characterized in their affinities for ADPbound actin monomers by Kd of 20 nM, 80 to 100 nM, 0.17 M, and0.4 M, respectively [reviewed in (42)]. The two ADP-bound actin
subunits, from the actin/twinfilin/CP complex, may then be converted to adenosine triphosphate (ATP)–bound actin by the actionsof CAP, due to the high affinity of CAP for ADP-bound actin(20, 21, 33, 43). Subsequently, the ATP-bound actin monomers will
be released to profilin, and possibly to thymosin-4, due to theirsuperior affinities for ATP-bound actin relative to twinfilin, CAP,
and cofilin, 0.1, 0.1 to 4.0, 0.5, 1.9, and 6 M (16, 23, 42), respectively.
This will replenish the polymerization-competent pool of actinmonomers (42). Because CAP also binds to the pointed ends ofactin filaments (33), association of the actin/twinfilin/CP complex
with the pointed end bound CAP may provide a mechanism to spatially separate the twinfilin/CP complex away from elongating actin
filament barbed ends close to the membrane.

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The fate of the twinfilin/CP complex may then be several-fold.
First, the twinfilin/CP complex may encounter a free barbed end ofa filament and undergo a further round of transient binding followedby dissociation of the terminal actin protomers. Thus, the twinfilin/
CP complex may depolymerize the filament in two subunit cycles,
maintaining transient capping while depolymerizing the filament.
Second, twinfilin may competitively dissociate from CP in favor of
CP-capped filaments due to the superior affinity for filament ends
relative to CP alone, Kd of 13 and 50 nM, respectively (17, 27). Thismechanism assumes that twinfilin affinity for the CP-capped filament is also high; however, the value is unknown and possibly difficult to measure due to the uncapping mechanism. Last, the twinfilin/
CP complex may be dissociated at the membrane by PIP2 or bycompetition with CPI proteins, such as CARMIL (27, 37).
The structure of the actin/twinfilin/CP complex suggests that asecond possible mechanism of filament uncapping may operate in
the presence of V-1. In this mechanism, the twinfilin/CP complex“wobbles” (44) at the barbed end of a filament, remaining attachedthrough the high-affinity actin-binding site on twinfilin D2, while
twinfilin D1 dissociates from actin (Fig. 6, A and C) (16). The extended portion of the twinfilin tail (residues 316 to 322, between theend of D2 and the first tail residue with significant contact with CP),
which we hypothesize is under tension, may aid the wobble process.
There is homology between twinfilin and CARMIL in this region(317-HAHKQSFAKP-326 in twinfilin-1 and 981-KLEHFTKLRP-990 in
CARMIL1) (27), which may allow the twinfilin tail to associate
more intimately with CP in a manner similar to the CARMIL CPI
interaction with CP (Fig. 3A) (34). Association of this region of
twinfilin with CP would hold CP away from the actin subunit, towhich it was previously bound via the -tentacle, thus freeing the
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27 January 2021

actin-binding site on the -tentacle to bind to V-1, and preventing
the -tentacle from reassociating with actin. The partially boundcomplex would then dissociate as twinfilin/CP/V-1 bound to a single actin, with the high-affinity twinfilin D2 inducing a G-actin–like
conformation in the actin subunit to destabilize its association withthe filament (16). Flow-cell total internal reflection fluorescencemicroscopy observations of CP uncapping by twinfilin detected enhanced uncapping in the presence of V-1 (28). This suggests that
V-1 may play an active role in twinfilin-aided uncapping as hypothesized in this mechanism (Fig. 6, A and C).
In summary, the actin/twinfilin/CP complex structure clarifies
the possible mechanisms by which twinfilin operates as a diffusing
uncapping agent. The majority of the in vitro observations outlined
above can be rationalized in terms of twinfilin’s proposed role intransforming CP from a strong capping agent into a transiently capping depolymerization complex, which aids in the recycling of actinmonomers and is sensitive to the presence of V-1. In the bulk cytoplasm, the outcome of twinfilin’s uncapping activities is likely to be
actin depolymerization while maintaining transient capping. Thisrole contrasts starkly to the CPI-containing proteins, which are target-­bound uncapping agents that are hypothesized to engender actin
polymerization for membrane remodeling (45, 46). The location of
uncapping, in either actin assembly- or disassembly-rich regions ofa cell, will determine whether filament polymerization or depolymerization will be the product of the uncapping. Any twinfilin-induced
uncapping at a membrane may therefore result in the opposite activity, actin polymerization. Because twinfilin/CP-bound filaments,
relative to CP-bound filaments, have enhanced barbed-end filament dynamics, we propose that cells are able to differentially regulate these actin filament barbed-end binding states to dictate the7 of 10

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Fig. 6. Models of the filament uncapping. (A) Cartoon comparison of CP and CP/twinfilin association at the barbed end of a filament. (B) Twinfilin-aided uncapping is a
result of twinfilin ADF-H domains inducing G-actin–like conformations in the terminal two actin protomers, weakening actin:actin interactions in the filament, leading to
the dissociation of the complex. (C) Twinfilin/V-1–aided uncapping requires space for V-1 to reach its binding site on CP via a wobble state of the twinfilin-bound complex.
Once V-1 is bound, CP is unable to reassociate with actin. In vitro effects of twinfilin alone on actin filaments and comparisons of uncapping models in the absence of
twinfilin are shown in fig. S9.

SCIENCE ADVANCES | RESEARCH ARTICLE
chromatography with a Superdex 75 pg column (GE Healthcare)equilibrated with 50 mM tris-HCl (pH 9) and 150 mM NaCl.
Human V-1 (pGEX-6P-3 vector) was expressed as a glutathione
S-transferase (GST) fusion E. coli BL21 Star (DE3) and affinity-­
purified on an ÄKTAxpress system (GE Healthcare) by loading cleared
lysate onto 1 ml of GSTrap FF Column (GE Healthcare) equilibrated
with 50 mM tris-HCl (pH 7.4), 150 mM NaCl, and 1 mM dithiothreitol (DTT). GST tag was removed by on-column digestion
with human rhinovirus 3C protease overnight followed by further
purification by gel filtration chromatography. CARMIL CP–binding
region (CBR) was expressed and purified as previously described (34).

MATERIALS AND METHODS

Preparation of the actin, twinfilin, and CP complex
Rabbit skeletal muscle actin was purified from skeletal muscle acetone powder (Pel-Freez) (53, 54). The protein was subjected to a
final SEC using HiLoad Superdex 200 on an ÄKTA Prime system.
The purity of G-actin was assessed by SDS-PAGE, and concentration was determined by measuring OD at 290 nm. The protein
complex was prepared by mixing human twinfilin-1, mouse CP 12,
and rabbit actin in buffer A [2 mM tris-HCl (pH 7.5), 0.2 mM ATP,
0.5 mM DTT, 1 mM NaN3, and 0.1 mM CaCl2] at a molar ratio of
1:1:2.5. The mixture was incubated on ice for 10 min to allow complex formation and then purified by SEC (HiLoad 16/60 Superdex
200 preequilibrated with buffer A) on an ÄKTAxpress system (GEHealthcare). Fractions corresponding to ultraviolet absorption peakwere analyzed by SDS-PAGE gel to verify complex formation.

Protein expression and purification
The gene sequences encoding full-length human twinfilin-1 andprofilin-1 were codon-optimized for Escherichia coli, synthesized(GenScript), and cloned into a pSY5 vector that includes anN-­terminal eight-histidine tag followed by a human rhinovirus 3C
protease cleavage site (51). The mouse full-length CP (1/2) construct was provided by P. Lappalainen (University of Helsinki, Finland).
The construct was in the pRSFDuet-1 vector designed for coexpression of two interacting target proteins and contains an N-terminal
six-histidine tag on the -subunit (52).
All constructs encoding proteins of interest were transformed
and recombinantly expressed in phage-resistant E. coli strain BL21
Star (DE3) (New England Biolabs). Fresh LB medium supplemented with either ampicillin (pSY5) or kanamycin (pRSFDuet-1)
was inoculated with respective overnight cultures and shaken at
37°C until the cell density reached OD 600 (optical density at
600 nm) ~ 0.6. Cells were induced for protein expression with
0.25 mM isopropyl-­-d-thiogalactopyranoside at 16°C overnight.
Cells were harvested via centrifugation at 4000g at 4°C for 1 hour,
and the pellets were resuspended in 50 ml of His-binding buffer[50 mM tris-HCl (pH 7.5), 500 mM NaCl, 20 mM imidazole, andone protease inhibitor tablet]. The cells were lysed by sonication
with a Vibra-Cell ultrasonic processor and clarified by centrifugation at 19,000 rpm in SS-34 rotor using an RC 5C Plus centrifuge(Sorvall) at 4°C for 1 hour followed by filtration through a 0.45-m
Minisart syringe filter (Sartorius). The proteins were purified on anÄKTAxpress system (GE Healthcare) by affinity chromatographyusing 5 ml of HisTrap FF column with or without (CP) on-column
cleavage of the His-tag with human rhinovirus 3C protease. Proteins
purified with the His-tag were eluted in buffer containing 50 mM
tris-HCl (pH 7.5), 500 mM NaCl, and 500 mM imidazole. The eluted
proteins were concentrated to 5 ml and subjected to size exclusion
chromatography (SEC) in a Superdex 75 pg column (GE Healthcare)preequilibrated with buffer containing 50 mM tris-HCl (pH 7.5)
and 150 mM NaCl. All purified proteins were verified by SDS–­
polyacrylamide gel electrophoresis (PAGE) before being snap-­frozen and stored at −80°C.
MBP-tagged twinfilin tail constituting residues Gln321-Asp350 was
polymerase chain reaction (PCR)–amplified from full-length human
twinfilin-1 expression plasmid and inserted into pSY7 vector, which
incorporates a cleavable N-terminal histidine and MBP tag. MBP
control was prepared by expressing pSY7 vector alone in BL21(DE3)
E. coli. Both MBP-tagged twinfilin tail and MBP were purified byaffinity chromatography using 1 ml of HisTrap FF column with on-­column digestion with human rhinovirus 3C protease to remove
the histidine tag. The proteins were further purified by gel filtration
Mwangangi et al., Sci. Adv. 2021; 7 : eabd5271

27 January 2021

Crystallization and crystal optimization
The fractions corresponding to the peak from SEC were pooled together, concentrated with Vivaspin 20 MWCO 10,000 concentrator(Sartorius) to approximately 20 mg/ml, and subjected to commercial crystallization screens. Crystallization screens were set up as
sitting drops in three-drop Intelli-Plate 96 (Hampton Research) in
2:1, 1:1, and 1:2 ratios consisting of protein solution and precipitantand stored in room temperature (25°C). Crystal hits were observedin a number of conditions in the JBScreen Classic HTS I screen after
3 days. One condition [12% polyethylene glycol (PEG) 8000, 10%
glycerol, and 500 mM potassium chloride] produced the best shaped
crystals composed of thin rods. A seed bead kit (Hampton Research)
was used to generate seed stocks of protein crystals for further optimization. Crystals were set up using the hanging-drop vapor diffusion method in 2-l drops of 1:1 ratio consisting of protein solution
and seed stock in the crystallization condition (12% PEG 8000, 10%
glycerol, and 500 mM potassium chloride).
Crystal data processing and structure determination
Harvested crystals were soaked in 25% glycerol before being carefully fished with cryoloops and flash-frozen in liquid nitrogen for
subsequent data collection at the National Synchrotron Radiation
Research Center, Taiwan. Indexing, scaling, and merging of datasets were carried out in HKL2000 (55). The crystal structure twinfilin/
CP/actin complex was solved at a resolution of 3.2 Å by molecularreplacement using Phaser (56), sequentially searching for one copy ofmouse twinfilin-1 D2 (3DAW) (19) and one copy of chicken CP hetero­dimer (3AA7) (35) followed by a second copy of mouse twinfilin-1
D2 (3DAW) (19). The resultant model was rebuilt by hand, and thestructure was subjected to several rounds of refinement using Phenix(56) and further manual rebuilding in COOT (57). The crystal datacollection and structure refinement statistics are presented in table S1.
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lifetimes of individual actin filaments. The proposed mechanism ofdepolymerization while maintaining transient filament capping
adds to the evidence that actin filament ends are highly controlled
and are rarely free in mammalian cells. During depolymerization,
filament barbed ends are controlled by twinfilin/CP and filament
pointed ends by CAP (33). During polymerization modes, the filament barbed ends can be regulated by formins (47), or alternatively
by the VASP family proteins (48), following either ARP2/3 nucleation or filament uncapping at the membrane (34, 40, 49, 50). In
each case, the filament end is protected.

SCIENCE ADVANCES | RESEARCH ARTICLE

SUPPLEMENTARY MATERIALSSupplementary material for this article is available at http://advances.sciencemag.org/cgi/
content/full/7/5/eabd5271/DC1

REFERENCES AND NOTES

1. O. L. Mooren, B. J. Galletta, J. A. Cooper, Roles for actin assembly in endocytosis. Annu. Rev.
Biochem. 81, 661–686 (2012).
2. T. D. Pollard, J. A. Cooper, Actin, a central player in cell shape and movement. Science 326,
1208–1212 (2009).
3. J. W. Rudy, Actin dynamics and the evolution of the memory trace. Brain Res. 1621, 17–28(2015).
4. T. D. Pollard, Actin and actin-binding proteins. Cold Spring Harb. Perspect. Biol. 8, a018226(2016).
5. A. D. Siripala, M. D. Welch, SnapShot: Actin regulators I. Cell 128, 626 (2007).
6. M. Poukkula, E. Kremneva, M. Serlachius, P. Lappalainen, Actin-depolymerizing factor
homology domain: A conserved fold performing diverse roles in cytoskeletal dynamics.
Cytoskeleton (Hoboken) 68, 471–490 (2011).
7. C. Akıl, L. T. Tran, M. Orhant-Prioux, Y. Baskaran, E. Manser, L. Blanchoin, R. C. Robinson,
Insights into the evolution of regulated actin dynamics via characterization of primitive
gelsolin/cofilin proteins from Asgard archaea. Proc. Natl. Acad. Sci. U.S.A. 117,
19904–19913 (2020).
8. P. W. Gunning, U. Ghoshdastider, S. Whitaker, D. Popp, R. C. Robinson, The evolutionof compositionally and functionally distinct actin filaments. J. Cell Sci. 128, 2009–2019 (2015).
9. C. Akıl, Y. Kitaoku, L. T. Tran, D. Liebl, H. Choe, D. Muengsaen, W. Suginta, A. Schulte,
R. C. Robinson, Mythical origins of the actin cytoskeleton. Curr. Opin. Cell Biol. 68, 55–63 (2020).
10. F. Rivero, F. Cvrcková, Origins and evolution of the actin cytoskeleton. Adv. Exp. Med. Biol.
607, 97–110 (2007).
11. B. L. Goode, D. G. Drubin, P. Lappalainen, Regulation of the cortical actin cytoskeleton
in budding yeast by twinfilin, a ubiquitous actin monomer-sequestering protein. J. Cell Biol.
142, 723–733 (1998).
12. S. Ono, Mechanism of depolymerization and severing of actin filaments and its
significance in cytoskeletal dynamics. Int. Rev. Cytol. 258, 1–82 (2007).
13. W. A. Elam, H. Kang, E. M. De la Cruz, Biophysics of actin filament severing by cofilin. FEBS
Lett. 587, 1215–1219 (2013).

Mwangangi et al., Sci. Adv. 2021; 7 : eabd5271

27 January 2021

14. M. Vartiainen, P. J. Ojala, P. Auvinen, J. Peranen, P. Lappalainen, Mouse A6/twinfilin isan actin monomer-binding protein that localizes to the regions of rapid actin dynamics.
Mol. Cell. Biol. 20, 1772–1783 (2000).
15. D. Wang, L. Zhang, G. Zhao, G. Wahlstrom, T. I. Heino, J. Chen, Y. Q. Zhang, Drosophila
twinfilin is required for cell migration and synaptic endocytosis. J. Cell Sci. 123,
1546–1556 (2010).
16. P. J. Ojala, V. O. Paavilainen, M. K. Vartiainen, R. Tuma, A. G. Weeds, P. Lappalainen, The
two ADF-H domains of twinfilin play functionally distinct roles in interactions with actinmonomers. Mol. Biol. Cell 13, 3811–3821 (2002).
17. E. Helfer, E. M. Nevalainen, P. Naumanen, S. Romero, D. Didry, D. Pantaloni,
P. Lappalainen, M. F. Carlier, Mammalian twinfilin sequesters ADP-G-actin and caps
filament barbed ends: Implications in motility. EMBO J. 25, 1184–1195 (2006).
18. V. O. Paavilainen, M. Hellman, E. Helfer, M. Bovellan, A. Annila, M. F. Carlier, P. Permi,
P. Lappalainen, Structural basis and evolutionary origin of actin filament capping by
twinfilin. Proc. Natl. Acad. Sci. U.S.A. 104, 3113–3118 (2007).
19. V. O. Paavilainen, E. Oksanen, A. Goldman, P. Lappalainen, Structure of the actindepolymerizing factor homology domain in complex with actin. J. Cell Biol. 182, 51–59(2008).
20. A. B. Johnston, A. Collins, B. L. Goode, High-speed depolymerization at actin filamentends jointly catalysed by Twinfilin and Srv2/CAP. Nat. Cell Biol. 17, 1504–1511 (2015).
21. D. M. Hilton, R. M. Aguilar, A. B. Johnston, B. L. Goode, Species-specific functionsof twinfilin in actin filament depolymerization. J. Mol. Biol. 430, 3323–3336 (2018).
22. J. B. Moseley, K. Okada, H. I. Balcer, D. R. Kovar, T. D. Pollard, B. L. Goode, Twinfilin isan actin-filament-severing protein and promotes rapid turnover of actin structuresin vivo. J. Cell Sci. 119, 1547–1557 (2006).
23. S. Falck, V. O. Paavilainen, M. A. Wear, J. G. Grossmann, J. A. Cooper, P. Lappalainen,
Biological role and structural mechanism of twinfilin-capping protein interaction. EMBO J.
23, 3010–3019 (2004).
24. M. A. Wear, A. Yamashita, K. Kim, Y. Maeda, J. A. Cooper, How capping protein binds
the barbed end of the actin filament. Curr. Biol. 13, 1531–1537 (2003).
25. J. A. Cooper, D. Sept, New insights into mechanism and regulation of actin capping
protein. Int. Rev. Cell Mol. Biol. 267, 183–206 (2008).
26. A. Yamashita, K. Maeda, Y. Maeda, Crystal structure of CapZ: Structural basis for actin
filament barbed end capping. EMBO J. 22, 1529–1538 (2003).
27. A. B. Johnston, D. M. Hilton, P. McConnell, B. Johnson, M. T. Harris, A. Simone,
G. K. Amarasinghe, J. A. Cooper, B. L. Goode, A novel mode of capping protein-regulationby twinfilin. eLife 7, e41313 (2018).
28. M. Hakala, H. Wioland, M. Tolonen, A. Jegou, G. Romet-Lemonne, P. Lappalainen,
Twinfilin uncaps filament barbed ends to promote turnover of lamellipodial actin
networks. bioRxiv 864769 [Preprint]. 4 December 2019. https://doi.org/10.1101/864769.
29. A. Narita, S. Takeda, A. Yamashita, Y. Maeda, Structural basis of actin filament capping at
the barbed-end: A cryo-electron microscopy study. EMBO J. 25, 5626–5633 (2006).
30. L. Urnavicius, C. K. Lau, M. M. Elshenawy, E. Morales-Rios, C. Motz, A. Yildiz, A. P. Carter,
Cryo-EM shows how dynactin recruits two dyneins for faster movement. Nature 554,
202–206 (2018).
31. T. Oda, M. Iwasa, T. Aihara, Y. Maeda, A. Narita, The nature of the globular- to fibrous-actin
transition. Nature 457, 441–445 (2009).
32. S. Z. Chou, T. D. Pollard, Mechanism of actin polymerization revealed by cryo-EM
structures of actin filaments with three different bound nucleotides. Proc. Natl. Acad. Sci.
U.S.A. 116, 4265–4274 (2019).
33. T. Kotila, H. Wioland, G. Enkavi, K. Kogan, I. Vattulainen, A. Jégou, G. Romet-Lemonne,
P. Lappalainen, Mechanism of synergistic actin filament pointed end depolymerization
by cyclase-associated protein and cofilin. Nat. Commun. 10, 5320 (2019).
34. M. Hernandez-Valladares, T. Kim, B. Kannan, A. Tung, A. H. Aguda, M. Larsson,
J. A. Cooper, R. C. Robinson, Structural characterization of a capping protein interaction
motif defines a family of actin filament regulators. Nat. Struct. Mol. Biol. 17, 497–503(2010).
35. S. Takeda, S. Minakata, R. Koike, I. Kawahata, A. Narita, M. Kitazawa, M. Ota, T. Yamakuni,
Y. Maeda, Y. Nitanai, Two distinct mechanisms for actin capping protein regulation—
Steric and allosteric inhibition. PLOS Biol. 8, e1000416 (2010).
36. T. Kim, G. E. Ravilious, D. Sept, J. A. Cooper, Mechanism for CARMIL protein inhibitionof heterodimeric actin-capping protein. J. Biol. Chem. 287, 15251–15262 (2012).
37. S. Palmgren, P. J. Ojala, M. A. Wear, J. A. Cooper, P. Lappalainen, Interactions with PIP2,
ADP-actin monomers, and capping protein regulate the activity and localization of yeast
twinfilin. J. Cell Biol. 155, 251–260 (2001).
38. S. Palmgren, M. Vartiainen, P. Lappalainen, Twinfilin, a molecular mailman for actinmonomers. J. Cell Sci. 115, 881–886 (2002).
39. K. Kim, A. Yamashita, M. A. Wear, Y. Maéda, J. A. Cooper, Capping protein binding to actin
in yeast: Biochemical mechanism and physiological relevance. J. Cell Biol. 164, 567–580(2004).

9 of 10

Downloaded from http://advances.sciencemag.org/ on April 8, 2021

Pyrene-actin polymerization assays
Pyrene-actin polymerization assays to monitor filament uncapping
were performed in 100-l reactions containing 2 M rabbit skeletal
muscle G-actin (10% pyrene–labeled), 0.5 M F-actin seeds, 2.8 M
profilin (to prevent pointed-end polymerization), 100 nM CP, andvariable concentrations of the test proteins. In all pyrene-actin polymerization assays, the final concentrations of twinfilin, CP, CARMIL,
V-1, MBP-tagged tail, and MBP were set at 1 M, 100 nM, 250 nM,
5 M, 10 M, and 10 M, respectively. The components were mixed
in buffer A [2 mM tris-HCl (pH 7.5), 0.2 mM ATP, 0.5 mM DTT,
1 mM NaN3, and 0.1 mM CaCl2] to a final volume of 70 l in a black
flat-bottom 96-well plate (Corning).
To prepare actin filament seed stocks, 5 M G-actin was polymerized for 1 hour at room temperature, after which the filaments
were mechanically sheared by repeatedly passing the F-actin solution through a 0.7-mm-diameter needle for 1 min. Thereafter, 10 lof the F-actin seed stock was mixed with 10 l of 1 M CP and left
for 5 min to allow barbed-end capping of the F-actin seeds. As acontrol, the F-actin seed stock was mixed with buffer A, without
CP. Then, 20 l of this precapped F-seed stock, or control, was added to pyrene-actin polymerization mixture and actin polymerization initiated by addition of 10 l of 10× KMEI buffer (500 mM KCl,
10 mM MgCl2, 10 mM EGTA, and 100 mM imidazole, pH 7.4) in a
total reaction volume of 100 l. The pyrene fluorescence intensities
were monitored immediately on a Safire2 fluorimeter plate reader(Tecan) with excitation and emission wavelengths set at 365 and407 nm, respectively.

SCIENCE ADVANCES | RESEARCH ARTICLE

Mwangangi et al., Sci. Adv. 2021; 7 : eabd5271

27 January 2021

54. J. A. Spudich, S. Watt, The regulation of rabbit skeletal muscle contraction. I. Biochemical
studies of the interaction of the tropomyosin-troponin complex with actin
and the proteolytic fragments of myosin. J. Biol. Chem. 246, 4866–4871 (1971).
55. Z. Otwinowski, W. Minor, Processing of X-ray diffraction data collected in oscillation
mode. Methods Enzymol. 276, 307–326 (1997).
56. P. D. Adams, P. V. Afonine, G. Bunkoczi, V. B. Chen, I. W. Davis, N. Echols, J. J. Headd,
L. W. Hung, G. J. Kapral, R. W. Grosse-Kunstleve, A. J. McCoy, N. W. Moriarty, R. Oeffner,
R. J. Read, D. C. Richardson, J. S. Richardson, T. C. Terwilliger, P. H. Zwart, PHENIX:
A comprehensive Python-based system for macromolecular structure solution.
Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 (2010).
57. P. Emsley, B. Lohkamp, W. G. Scott, K. Cowtan, Features and development of Coot.
Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).
58. L. Holm, L. M. Laakso, Dali server update. Nucleic Acids Res. 44, W351–W355 (2016).
59. S. Nag, M. Larsson, R. C. Robinson, L. D. Burtnick, Gelsolin: The tail of a molecular gymnast.
Cytoskeleton (Hoboken) 70, 360–384 (2013).
Acknowledgments: We thank B. Xue for technical advice, S. Takeda and B. Xue for valuable
discussions, P. Lappalainen (University of Helsinki, Finland) and J. Cooper (Washington
University School of Medicine, USA) for constructs, and the experimental facility and thetechnical services provided by the Synchrotron Radiation Protein Crystallography Facility of
the National Core Facility Program for Biotechnology, Ministry of Science and Technology and
the National Synchrotron Radiation Research Center, a national user facility supported by the
Ministry of Science and Technology, Taiwan. Funding: This work was supported by Biomedical
Research Council of A*STAR under the Singapore International Graduate Award (SINGA)
scholarship, by NMRC Grant OFIRG/027/2016, and by Human Frontiers Science Program award
RGP0028/2018. Author contributions: D.M.M. and R.C.R. conceived experiments, analyzed
data, and wrote the paper. D.M.M. performed experiments. R.C.R. and E.M. supervised thework. Competing interests: The authors declare that they have no competing interests. Dataand materials availability: The atomic coordinates and structure factors have been
deposited in the Protein Data Bank (PDB) under the accession code 7CCC. All other data are
available from the corresponding author upon reasonable request.
Submitted 26 June 2020
Accepted 8 December 2020
Published 27 January 2021
10.1126/sciadv.abd5271
Citation: D. M. Mwangangi, E. Manser, R. C. Robinson, The structure of the actin filament
uncapping complex mediated by twinfilin. Sci. Adv. 7, eabd5271 (2021).

10 of 10

Downloaded from http://advances.sciencemag.org/ on April 8, 2021

40. D. A. Schafer, P. B. Jennings, J. A. Cooper, Dynamics of capping protein and actin
assembly in vitro: Uncapping barbed ends by polyphosphoinositides. J. Cell Biol. 135,
169–179 (1996).
41. K. Tanaka, S. Takeda, K. Mitsuoka, T. Oda, C. Kimura-Sakiyama, Y. Maeda, A. Narita,
Structural basis for cofilin binding and actin filament disassembly. Nat. Commun. 9, 1860(2018).
42. B. Xue, R. C. Robinson, Guardians of the actin monomer. Eur. J. Cell Biol. 92, 316–332 (2013).
43. T. Kotila, K. Kogan, G. Enkavi, S. Guo, I. Vattulainen, B. L. Goode, P. Lappalainen, Structural
basis of actin monomer re-charging by cyclase-associated protein. Nat. Commun. 9, 1892
(2018).
44. N. Bhattacharya, S. Ghosh, D. Sept, J. A. Cooper, Binding of myotrophin/V-1 to actincapping protein: Implications for how capping protein binds to the filament barbed end.
J. Biol. Chem. 281, 31021–31030 (2006).
45. A. I. Fokin, V. David, K. Oguievetskaia, E. Derivery, C. E. Stone, L. Cao, N. Rocques,
N. Molinie, V. Henriot, M. Aumont-Nicaise, M.-V. Hinckelmann, F. Saudou, C. Le Clainche,
A. P. Carter, G. Romet-Lemonne, A. M. Gautreau, The Arp1/11 minifilament of dynactin
primes the endosomal Arp2/3 complex. bioRxiv 2020.05.29.123372 [Preprint]. 29 May 2020.
https://doi.org/10.1101/2020.05.29.123372.
46. M. H. Lanier, P. McConnell, J. A. Cooper, Cell migration and invadopodia formation
require a membrane-binding domain of CARMIL2. J. Biol. Chem. 291, 1076–1091 (2016).
47. S. H. Zigmond, M. Evangelista, C. Boone, C. Yang, A. C. Dar, F. Sicheri, J. Forkey, M. Pring,Formin leaky cap allows elongation in the presence of tight capping proteins. Curr. Biol.
13, 1820–1823 (2003).
48. S. D. Hansen, R. D. Mullins, VASP is a processive actin polymerase that requires
monomeric actin for barbed end association. J. Cell Biol. 191, 571–584 (2010).
49. M. D. Welch, A. Iwamatsu, T. J. Mitchison, Actin polymerization is induced by Arp2/3
protein complex at the surface of Listeria monocytogenes. Nature 385, 265–269(1997).
50. P. A. Janmey, C. Chaponnier, S. E. Lind, K. S. Zaner, T. P. Stossel, H. L. Yin, Interactionsof gelsolin and gelsolin-actin complexes with actin. Effects of calcium on actin nucleation,
filament severing, and end blocking. Biochemistry 24, 3714–3723 (1985).
51. S. Nag, Q. Ma, H. Wang, S. Chumnarnsilpa, W. L. Lee, M. Larsson, B. Kannan,
M. Hernandez-Valladares, L. D. Burtnick, R. C. Robinson, Ca2+ binding by domain 2 playsa critical role in the activation and stabilization of gelsolin. Proc. Natl. Acad. Sci. U.S.A. 106,
13713–13718 (2009).
52. Y. Soeno, H. Abe, S. Kimura, K. Maruyama, T. Obinata, Generation of functional
beta-actinin (CapZ) in an E. coli expression system. J. Muscle Res. Cell Motil. 19, 639–646(1998).
53. H. Wang, R. C. Robinson, L. D. Burtnick, The structure of native G-actin. Cytoskeleton(Hoboken) 67, 456–465 (2010).

The structure of the actin filament uncapping complex mediated by twinfilin
Dennis M. Mwangangi, Edward Manser and Robert C. Robinson

Sci Adv 7 (5), eabd5271.
DOI: 10.1126/sciadv.abd5271originally published online January 27, 2021

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