Biochemical Implications of Three-Dimensional Model of Monomeric Actin Bound to Magnesium-Chelated ATP

Structure
Article
Biochemical Implications of a Three-Dimensional
Model of Monomeric Actin Bound to
Magnesium-Chelated ATP
Keiji Takamoto,1,2,* J.K. Amisha Kamal,1,2 and Mark R. Chance1,2
1Case Center for Proteomics, Case Western Reserve University, 10090 Euclid Avenue, Cleveland, OH 44106, USA
2Lab address: http://casemed.case.edu/proteomics/
*Correspondence: keiji.takamoto@case.edu
DOI 10.1016/j.str.2006.11.005
SUMMARY
Actin structure is of intense interest in biology
due to its importance in cell function and
motility mediated by the spatial and temporal
regulation of actin monomer-filament intercon-versions
in a wide range of developmental and
disease states. Despite this interest, the struc-ture
of many functionally important actin forms
has eluded high-resolution analysis. Due to the
propensity of actin monomers to assemble into
filaments structural analysis of Mg-bound actin
monomers has proven difficult, whereas high-resolution
structures of actin with a diverse ar-ray
of ligands that preclude polymerization have
been quite successful. In this work, we provide
a high-resolution structural model of the Mg-
ATP-actin monomer using a combination of
computational methods and experimental foot-printing
data that we have previously published.
The key conclusion of this study is that the
structure of the nucleotide binding cleft defined
by subdomains 2 and 4 is essentially closed,
with specific contacts between two subdo-mains
predicted by the data.
INTRODUCTION
Actin is a ubiquitous and important protein in eukaryotes
and is extremely well conserved from yeast to man. Actin
binding of nucleotides and its interactions with other pro-teins
in the cell control the spatial and temporal assembly
and disassembly of the cytoskeletal network; the careful
regulation of this network has a profound influence on
cell motility (Paavilainen et al., 2004; Schoenenberger
et al., 2002; Wear et al., 2000; Winder, 2003). Even the
nature of the metal ion bound to the nucleotide with the
actin structure can profoundly alter the ability of actin
monomers to assemble into filaments. In spite of its bio-logical
importance, the structure of the Mg2+-ATP-bound
form of the actin monomer (called G-actin) is not known.
The determination of the high-resolution structure of
Mg2+-G-actin is hampered by the propensity of actin to
polymerize. Generation of crystals suitable for X-ray dif-fraction
as well as NMR studies typically requires higher
concentrations than the critical concentration for actin
polymerization for the Mg2+-nucleotide-bound forms of
the actin monomer. The high-resolution crystal structures
of G-actin, whose richness and depth are a nearly unique
resource for this investigation, never the less have either
Ca2+-nucleotide bound to actin or, if Mg2+-bound nucleo-tide
is used, actin is cocrystallized with ligands such as
DNase I (Kabsch et al., 1985, 1990; Suck et al., 1981), gel-solin
segment 1 (Mannherz et al., 1992; Vorobiev et al.,
2003), vitamin D binding protein (Otterbein et al., 2002;
Swamy et al., 2002; Verboven et al., 2003), or macrolides
(Allingham et al., 2004; Klenchin et al., 2003; Morton et al.,
2000; Reutzel et al., 2004; Yarmola et al., 2000). These li-gands
typically prevent polymerization. Although struc-tures
of Ca2+- and Mg2+-G-actin bound to ligands are
overall similar in many respects, significant differences in
structure and function between these species in the
absence of other bound ligands in solution have been
consistently reported, including biochemical/biophysical
analyses such as limited proteolysis (Chen et al., 1995;
Strzelecka-Golaszewska et al., 1993), fluorescence stud-ies
(Frieden and Patane, 1985; Moraczewska et al., 1999;
Selden et al., 1989; Valentin-Ranc and Carlier, 1991; Zim-merle
et al., 1987), and molecular dynamics simulations
(Wriggers and Schulten, 1997).
Recently, we have investigated the solution structure of
Mg2+-ATP-actin using hydroxyl-radical footprinting and
mass spectrometry (MS); these experiments have in-cluded
a comparison of Ca2+-ATP-actin in the presence
and absence of gelsolin segment 1 (GS1) (Guan et al.,
2003). The measured side-chain solvent accessibilities
of Ca2+-ATP-actin in the absence of GS1 are very similar
to that in the presence of GS1 overall and consistent with
the accessible surface area (ASA) calculated from solved
crystal structures. In contrast, Mg2+-ATP-actin in the
absence of GS1 is quite different in its side-chain surface
accessibilities, particularly in subdomains (SD) 2 and 4.
These differences are reversed in the presence of GS1,
indicating the structure of the Mg2+-ATP-actin/GS1 com-plex
in solution is consistent with crystal structure data.
Structure 15, 39–51, January 2007 ª2007 Elsevier Ltd All rights reserved 39

Hydroxyl-radical footprinting and MS have been proven
to be powerful tools for probing protein structure (Guan
et al., 2003; Rashidzadeh et al., 2003) and conformational
change (Kiselar et al., 2003a, 2003b), as well as for probing
protein-ligand (Gupta et al., 2004) and protein-protein
interactions (Guan et al., 2002, 2004; Liu et al., 2003). As
the experimental details of the technique have matured
and become reliable (Guan and Chance, 2005; Takamoto
and Chance, 2006), we and others have attempted to use
the data in conjunction with comparative modeling tech-niques
to provide unique structural models (Gupta et al.,
2004; Sharp et al., 2005). In this study, we extend our
previous work (Guan et al., 2003) with visualization of
our descriptive prediction by generating an atomic model
of the Mg2+-ATP-actin monomer structure using footprint-ing
and crystallographic data. The computational strategy
uses rigid-body rotations and translations of actin subdo-mains,
primarily guided by known subdomain rearrange-ments
from the range of actin crystallographic structures,
in order to generate a structure consistent with surface
accessibility data predicted by footprinting. In addition
to an atomic model of the Mg2+-ATP-actin monomer, we
also propose a specific mechanism for cleft closure
mediated by changes in metal-ion coordination.
RESULTS
Rigid-Body Rearrangements of Actin
Domain/Subdomains Derived from
an Actin Crystallographic ‘‘Database’’
The wealth of actin structures available in the literature
provides a nearly unique opportunity for examining the
range of conformations accessible to actin in its various
ligand-bound states. Actin is composed of two domains
that are termed the large and small domains, and each do-main
is composed of two subdomains (Figures 1A and
1B). Table 1 provides a ‘‘database’’ of structures that
encompass a number of relative arrangements of the var-ious
actin subdomains. These are the major structures
that are used in this paper to provide an analysis of the
range of motions of the actin subdomains. Our approach
in this section is to survey these structures and determine
patterns of subdomain motions.
Our previous footprinting data (Guan et al., 2003)
indicated that the large cleft (nucleotide binding cleft) be-tween
SD2 and SD4 is in a more closed configuration for
monomeric actin bound to Mg2+-ATP compared to the
Ca2+-ATP form. We surveyed the available crystal struc-tures
to ascertain the range of relative motions of the large
cleft. Most actin crystal structures (Table 1) are very
similar, with an overall backbone rmsd of 0.8 A°
for 13
structures (including Protein Data Bank ID codes 1RFQ-A
and 1RFQ-B, 1IJJ-A and 1IJJ-B, 1NM1, 1D4X, 1EQY-A,
1NWK, 1QZ6, 1YAG, 1MA9, 1AQK, and 1NLV). However,
there are some crystal structures that show significant de-viations
from the majority of solved structures. One of the
most prominent examples is crystal structure 1HLU (Chik
et al., 1996), which has profilin bound to the small cleft
Figure 1. Domain Rigid-Body Movements Observed in Actin
Crystal Structures
(A) The movement of the large domain relative to SD1 observed in crys-tal
structure 1HLU. The models shown in tin and silver are 1YAG and
1HLU, respectively. The movements of Ca atoms are indicated by
arrows. The colors of arrows indicate the size of movements ranging
from blue (small) to orange (large). The center of rotational movement
and normal vector are shown by the yellow sphere and long arrow.
(B) Schematic representation of large-domain movement.
(C) The movements of SD2 relative to subdomain 1. The white arrows
indicate deviations of the G63 Ca atom position from 1YAG.
between SD1 and SD3; this structure exhibits a ‘‘super-open’’
nucleotide cleft, with a large movement of the
ATP/cation complex within the cleft. This structure can be
related to other more ‘‘typical’’ structures through domain
movements by rotational/sheer transformation (Chik et al.,
1996; Page et al., 1998). A second example is 1RFQ
40 Structure 15, 39–51, January 2007 ª2007 Elsevier Ltd All rights reserved
Structure
Model of Mg2+-ATP-Actin Monomer

Table 1. List of Crystal Structures Used for Analyses
Protein Data Bank ID Code Species Cocrystal Cofactors Resolution Comments
Crystal Structures of Interest
1YAG Yeast GS1b Mg2+/ATP 1.90 ‘‘Template’’ molecule (with
1RFQ-Ba Rabbit LARc Mg2+/ATP 3.00 Closed large cleft with altered
1QZ5 Rabbit KABd Ca2+/ATP 1.45 Closed large cleft
1HLU Bovine Profilin Ca2+/ATP 2.65 Large cleft in ‘‘super-open’’
Crystal Structures with Complete D Loop
1J6Z Rabbit TMR Ca2+/ATP 1.54 TMR affects C-terminal structure?
1YVN Yeast GS1 Mg2+/ATP 2.10 V159N mutant of 1YAG
2BTF Bovine Profilin Sr2+/ATP 2.55 With Sr2+. b-actin
1ATN Rabbit DNase I Ca2+/ATP 2.80 D loop interacts with DNase I
1IJJ Rabbit LAR Mg2+/ATP 2.85 D loop that interacts with another
1LCU Rabbit LAR Ca2+/ATP 3.50
Crystal Structures with Partial D Loop
1D4X C. elegans GS1b Ca2+/ATP 1.75
1C0G Chimeric GS1b Ca2+/ATP 2.00 Chimeric, Q228K/T229A/A230Y/E360H
1MDU Chicken GS1b Ca2+/ATP 2.20 Actin-trimer
1C0F Chimeric GS1b Ca2+/ATP 2.40 Chimeric Dictyostelium/Tetrahymena
1DEJ Chimeric GSe Mg2+/ATP 2.40 Chimeric, Q228K/T229A/A230Y/
1H1V Human GSe Ca2+/ATP 3.00
Important crystal structures of monomeric actin. The first set is used for modeling process or strategy; the second set is actin struc-tures
with complete D-loop backbone coordinates; and the last set is a partial but relatively better coverage of D-loop backbone
coordinates. Except for 1RFQ, the interasymmetric unit interactions between their D loop and the other molecule are unknown.
1RFQ does not have contact with other molecules.
a 1RFQ-B, chain B of asymmetric unit in 1RFQ.
b Gelsolin segment 1.
c Latrunculin A.
d Kabiramide C.
e Gelsolin.
(Reutzel et al., 2004), which has two chains in the asym-metric
unit whose structures are quite different from
each other. Chain A exhibits a ‘‘standard’’ cleft structure
similar to those of the vast majority of solved structures.
However, chain B (1RFQ-B) is interesting as it shows
a ‘‘closed’’ form of the cleft, with very different geometries
for the residues inside the cleft. The closed form in chain B
of the crystal structure exemplifies a database entry that
qualitatively satisfies an important aspect of our footprint-ing
data, that is, it provides an example of how relative
subdomain motions can result in a closed nucleotide cleft.
Crystal structure 1QZ5 (Klenchin et al., 2003) shows a
closed large cleft as well. In this structure, the geometry in-side
the large cleft is almost identical to 1YAG (although it
has a Ca2+ ion instead of an Mg2+ ion). To achieve cleft
closure compared to canonical actin forms, SD2 moves
complete D loop)
side-chain geometries
state, b-actin
chain’s small cleft
A231K/S232E/E360H
toward SD4 without tilting (unlike 1RFQ-B), minimizing ste-ric
conflicts between the turn in SD2 (residues 62–65) and
the top of the helix in SD4 (residues 200–205). Although our
hydroxyl-radical footprinting results indicate more signifi-cant
closure of the cleft than observed in these structures,
these structures can serve as ‘‘qualitative’’ templates for
understanding the Mg2+ form of G-actin in solution. The
B factors of the atoms in SD2 of 1QZ5 are relatively high,
indicating the relative mobility of SD2. This suggests the
likelihood of high relative mobility of SD2 (not only the D
loop but also the SD2 core) in solution (see the Supple-mental
Data available with this article online).
Vector Analysis of Subdomain Relative Positions
The structure 1YAG is used as our standard structural
template having high-resolution (1.90 A°
) and well-defined
Structure

waters within the nucleotide binding pocket. The struc-tures
of Protein Data Bank ID codes 1HLU (Chik et al.,
1996), 1QZ5 (Klenchin et al., 2003), 1RFQ-B (Reutzel
et al., 2004), 1J6Z (Otterbein et al., 2001), 1ATN (Kabsch
et al., 1990), 1S22 (Allingham et al., 2004), 1EQY-A
(Mannherz et al., 1992), 1IJJ-A (Bubb et al., 2002), 1MA9
(Verboven et al., 2003), 1KXP (Otterbein et al., 2002),
and 2BTF (Schutt et al., 1993) were compared with 1YAG,
and differences between Ca/backbone coordinates were
visualized using Tcl scripts for visual molecular dynamics
(VMD). The script also calculates the average center
position and rotational normal vector of movement of
Ca/backbone coordinates.
Figure 1A shows the relative movements (vector) of the
large domain with respect to SD1 in the comparison of
1YAG (the standard) and 1HLU (most open nucleotide
cleft structure). The arrows indicate the Ca movements
(arrows from coordinates in 1YAG to 1HLU). As the move-ments
are relative, we chose to show the large-domain
movement relative to SD1 in this representation because
SD2 also shows significant movement against SD1. As
seen in Figure 1B, this movement is 8.9 rotational (hinge)
movement that has a center position in the junction be-tween
SD1 and SD3 (around Q137-A138). The axis of ro-tation
goes from the front to the back side of the molecule
(where the front side is defined as the surface of the actin
molecule that exposes ATP). Figure 1B illustrates this
movement (the rotational angle is exaggerated for clarity).
A similar movement is reported in previous analyses and
refinement of F-actin fiber X-ray data including alterations
of the nucleotide binding pocket (Lorenz et al., 1993; Tirion
et al., 1995).
The SD2 subdomain is extremely mobile even in crystal
structures. Figure 1C compares 11 SD2 structures with
that of 1YAG; SD2 is observed in many orientations rela-tive
to SD1. The directions of rotation of these crystal
structures (relative to 1YAG) are shown in Figure 1C by
arrows. All of these differences can be mediated by
hinge movements having pivot points near P32-S33 and
Y69-P70.
Intracleft Interactions
There are a number of interactions between the ATP/
cation complex and the protein within this cleft that are
important for understanding large-cleft structure and the
possible mechanisms of the structural changes. The sub-domains
are not strongly connected by hydrogen-bond
networks or other forms of interactions; the majority of
the interactions between subdomains are mediated
through a hydrogen-bond network with the ATP molecule
and with waters in the pocket of the cleft. SD1 has interac-tions
with phosphate oxygen atoms of ATP through resi-dues
S14, G15, L/M16, and K18 (Figure 2A). Connections
between SD1 and SD2 are mediated only by the hydrogen
bonds between S14 and G74 (Figure 2B) (Chen et al.,
1995; Kabsch et al., 1990; Schu¨ ler, 2001). This link is
severed in 1RFQ-B due to subdomain rearrangements.
Although the metal ion is located near the interface of sub-domains
1 and 2, it only interacts with SD1 by hydrogen
Figure 2. Important Interactions Involving the ATP Molecule
Brown and silver colors are 1YAG and 1FRQ, respectively. Hydrogen
bonds and coordination bonds are shown by broken lines.
(A) N-terminal b strand and ATP-metal interactions.
(B) Hydrogen bonds that connect subdomains 1 and 2 and ATP.
(C) Interactions with the Mg2+ ion. Water molecules observed in 1YAG
are shown as blue balls.
bonds through waters (no inner-sphere coordination)
and no interaction is formed with SD2 residues.
The interactions between SD3 and SD4 are also mainly
mediated through ATP. In addition, there is a poorly
packed hydrophobic cluster at the interface of the two
Structure

Table 2. Probe Residue Solvent Accessibilities from Crystal Structures and Experimentally Determined
Hydroxyl-Radical Reactivities
Crystal Structures
1ATN 1YAG 1RFQB 1RDW Footprinting Data
Ca Mg Mg Mg Solution G-Actin
DNase I GS1 LAR LAR Ca Mg
subdomains. The adenosine moiety of ATP is located in
the pocket between the two subdomains, with hydrogen
bond and van der Waals contacts with surrounding resi-dues.
Thus, SD3, SD4, and ATP form an interdependent
set of interactions.
Metal-Ion Coordination
In most nonactin protein structures with ATP bound, mag-nesium
ion coordinates to at least one residue from the
protein (14 structures out of 16 nonredundant structures
examined; data not shown; Dudev et al., 1999). In the
case of actin, the magnesium ion (or calcium ion) forms in-ner-
sphere coordination bonds with phosphate oxygens
and water molecules, but not side-chain oxygen atoms,
with a coordination number of 6 (or 7 for Ca2+). A notice-able
difference between 1YAG (or other structures) and
1RFQ-B is that residue atoms Q137:OE1 and D11:OD1,
OD2 are closer to the Mg2+ ion in 1RFQ-B (3.0, 3.5, and
4.0 A °
) compared to 1YAG (4.4, 4.3, and 4.2 A °
, respec-tively).
Figure 2C shows the difference in the geometries
surrounding the magnesium ion. Unfortunately for 1RFQ,
at a resolution of 3.0 A °
, ordered water molecules are not
observed within the nucleotide binding cleft. However,
some water molecules observed in 1YAG must be radi-cally
reorganized in 1RFQ-B, as these water molecules
would clash with side-chain oxygen atoms (Figure 2C,
pink side-chain oxygen atoms overlapping with blue water
oxygen atoms). Based on these data, we used the geom-etry
inside the large cleft from 1RFQ-B as a template for
our structural modeling of the nucleotide binding cleft.
Comparison of Crystallographic Data and
Hydroxyl-Radical Footprinting Data
Table 2 summarizes the calculated ASAs and rate con-stants
of modification of actin peptides analyzed by
hydroxyl-radical footprinting (Guan et al., 2003).
The hydroxyl-radical footprinting data for Ca2+-ATP-G-actin
are generally consistent with the calculated ASA,
that is, solvent-accessible, reactive residues show oxida-tion
and inaccessible residues are not appreciably oxi-dized.
However, a number of probe residues in SD2 and
SD4 and at the C terminus (SD1) show decreased rates
of modification for Mg2+-ATP-G-actin where the solvent
accessibilities for the structure are similar to those for
Ca2+-ATP-G-actin. These sites that show protections
from oxidation for Mg- versus Ca-actin are illustrated in
Residue
Number Residue
Side-chain
ASA (A° 2)
Side-chain
ASA (A° 2)
Side-chain
ASA (A° 2)
Side-chain
ASA (A° 2)
Rate
constant
(s1)
Rate
constant
(s1)
Peptide
(Residue
Numbers)
21 Phe 34.32 23.87 37.13 34.39 0.65/0.70 0.32/0.43 19–28 11–23
44 Met 79.35 (16.31)a 137.63 (35.96)a N.D.b N.D.b 33 20 40–50
47 Met 104.58 (30.94)a 113.37 (24.81)a N.D.b N.D.b 33 20
53 Tyr 14.26 6.60 64.67c 55.88c 0.69 0.32 51–61
67 Leu 28.70 22.16 25.46 40.25 0.40 0.08 63–68
69 Tyr 80.46 98.57 68.54 66.81 1.1 0.18 58–72
200 Phe 0.36 3.61 10.79 5.50 1.10/0.83 0.26/0.24 197–206
196–207
201 Val 82.01 65.45 108.42 101.42 1.10/0.83 0.26/0.24
202 Thr 81.61 72.45 43.79 73.26 1.10/0.83 0.26/0.24
243 Pro 43.26 63.97 72.23 68.46 0.69/0.75 0.26/0.30 239–254
242–253
362 Tyr 12.59 9.05 15.18 11.75 0.72 0.38 362–372
367 Pro 45.75 40.74 39.49 41.88 0.72 0.38
371 His 27.79 26.53 34.71 25.89 0.72 0.38
374 Cys N.D. 3.53 49.40 24.29 8.5 3.6 363–375
375 Phe N.D. 157.72 77.56 44.38 N.D.d N.D.d
a Values in parentheses are ASAs for the sulfur atom in the side chain.
b These crystals lacking observed D-loop structures.
c These values are affected by the lack of a D loop (larger than expected).
d The rate constant in the peptide is dominated by C374 and thus could not be determined.
Structure

Figure 3. For example, probe residues in the D loop are
modestly protected from oxidative modifications (40%
reduction in rate constant), whereas residues 200–202
and 243 also experience strong reductions in modification
rate (75% and 60%, respectively). L67 and Y69 lo-cated
inside the large cleft (Figure 3) experience an 80%
reduction in modification rates. Consistent with these find-ings
are limited proteolysis data, where cleavage between
K68 and Y69 is almost completely suppressed in Mg2+-
G-actin compared to Ca2+-G-actin (Chen et al., 1995;
Strzelecka-Golaszewska et al., 1993). A closure of the
large cleft is consistent with these data.
Modeling Strategy: Overall Structural
Considerations
In the previous section, we analyzed possible relative
movements of domains/subdomains. In light of these
analyses, we can understand the cleft structures in each
case. DNase I binding prevents large-cleft closure by
prohibiting the movements of SD2 and the large domain
by interacting with both of them (Kabsch et al., 1990).
Latrunculin A binds between SD2 and SD4 (Bubb et al.,
2002; Reutzel et al., 2004); this prevents movements of
those subdomains. Gelsolin segment 1 (Mannherz et al.,
1992; McLaughlin et al., 1993) and vitamin D binding pro-tein
(Otterbein et al., 2002) prevent the domain movement
between SD1 and SD3 blocking corresponding large-domain
movements. Some macrolides (kabiramide C,
jaspisamide A, and ulapualide A; Allingham et al., 2004;
Klenchin et al., 2003) bind to the small cleft and block its
movement as well. This is summarized in Figure 4A. It is
important to note that macrolides are small enough to fit
into small or large clefts and do not seem to prevent poly-merization
by steric hindrance as opposed to actin binding
proteins that result in steric blockage. It appears that
these macrolides interfere with polymerization by prevent-ing
domain/subdomain motions.
The analysis of the various crystal structures and the
footprinting data (Guan et al., 2003) provides significant
clues about how the structure of Mg2+-G-actin differs
from that of Ca2+-G-actin. First, the footprinting data indi-cate
that the large cleft between SD2 and SD4 is almost
completely closed. Second, the analyses of the various
crystal structures indicate that movement of the SD2 core
is a rigid-body movement. The movement of SD2 toward
the large domain alone cannot explain the protections of
Figure 3. Analyses of Hydroxyl-Radical Reactivity and Struc-tures
The protection sites in Mg2+-G-actin (compared with Ca2+-G-actin) are
displayed with residues. Colors are coded as blue (strong protection)
to red (enhancement) by reactivity changes.
Figure 4. Schematic Representation of Domain/Subdomain
Movements
(A) The effects on subdomain movements by ligand binding.
Actin binding proteins (top) bind to actin and prevent polymerization,
possibly by steric hindrance or interference of domain movements.
Macrolides are small enough to fit into small or large clefts and interfere
with the domain movements. They may prevent polymerization by this
mechanism.
(B) The proposed mechanism for large-cleft closure. SD2 movement
alone cannot explain the complete closure of the large cleft (top right).
The partial closure by large-domain movement brings SD4 and SD2
close enough to allow interactions between the two subdomains, re-sulting
in complete cleft closure.
Structure

both L67 and Y69. Y69 is located deep within the large cleft
and cannot be buried by SD2 rigid-body movement. In ad-dition,
neither M44 nor M47 can reach SD4 to form pair-wise
interactions to protect P243 and M44/47. This sug-gests
that both the large domain and SD2 need to move
as rigid bodies relative to SD1 in order to explain protec-tions
observed in SD2 and SD4 (Figure 4B).
The hydrogen-bond network among phosphate, S14,
and G74 is the only one actually connecting SD2 to SD1
and is observed in almost all structures (Schu¨ ler, 2001).
This linkage is severed in the 1RFQ-B structure. We spec-ulate
that latrunculin A prevents the movement of SD2 fur-ther
from the observed structure although the geometry
inside the large cleft is changed. As a result, the hydrogen
bonds are broken. This hydrogen-bond linkage was main-tained
in the model structure. We constructed a series of
structures with combinations of different rotational angles
and normal vectors for SD2 in order to determine the best-fit
model for the hydroxyl-radical data.
Regions Excluded from Modeling
Although there is evidence for structural variation in the
C-terminal region (Y362 to F375 changes solvent accessi-bility
in hydroxyl-radical footprinting; Guan et al., 2003)
and other data (Frieden and Patane, 1985; Strzelecka-
Golaszewska et al., 1993; Valentin-Ranc and Carlier, 1991;
Zimmerle et al., 1987), we do not have clear evidence to
support the specific structural differences. Thus, we have
not modeled the structural differences in this region. F21
experiences an 50% reduction in modification rate in
Mg2+-G-actin compared to Ca2+-G-actin, but this change
cannot be explained with our modeling strategy. Our
speculation is that the C-terminal (residues 336–375)
and N-terminal (residues 1–33) regions interacting with
SD1 are also affected by the movements of SD2 (con-nected
to the N-terminal region) and the large domain
(connected to the C-terminal region) and mediate these
additional structural differences between the two forms.
Model of the Mg2+-ATP-Actin Monomer
In general, the hydroxyl-radical footprinting data are
very consistent with known crystal structures. Solvent-accessible
surface areas and rate constants of oxidation
on side chains in known structures are in good correlation
(Guan et al., 2004; Guan and Chance, 2005; Takamoto
and Chance, 2006; Xu and Chance, 2005). This is not sur-prising,
as the size of the hydroxyl radical is very close to
that of a water molecule. Although it is not easy to com-pare
rate constants among different side chains (such as
Leu and Phe), it is very quantitative and reliable for the
same residue in different conformational states. The site
of oxidation is determined by MS/MS analyses that are
well established and routine. With the use of different
proteases, protein sequence coverage by MS analysis is
usually 80%–90% and most of the probe residues can
be detected. In our previous study, we covered 90% of
the actin sequence with trypsin and Asp-N proteases.
Thus, it is important to note that we have used high-quality
data for our modeling.
Figure 5. The Model
(A) Overlaid structure of the model (silver) and template structure 1YAG
(brown). The residues that experience protections are displayed as
sticks and bubbles. Structures are aligned by subdomain 1.
(B) The difference between the model and template structure 1YAG.
The arrows indicate the differences of backbone atom positions. The
sphere and long arrow indicate the center of rotation and its normal
vectors for rigid-body movements (green for large domain and yellow
for subdomain 2).
Figure 5A shows the final model and the template struc-ture
for Mg2+-G-actin. The residues indicated to change
conformation in the footprinting experiments are shown
in stick-and-bubble format for both structures. The struc-tures
are aligned at SD1 in order to show both large-domain
and SD2 movements. The large domain rotates
toward the small domain and SD2 rotates to the front
(as defined previously) and toward the large domain.
Figure 5B show the backbone movement analyzed by
VMD/Tcl scripts. Figures 6A and 6B show a magnified view
of the SD2/SD4 interface in the model. In order to make
contact between D-loop residue M44 and the hydropho-bic
pocket in SD4 around protected residue P243, SD2
was moved toward the front side along with its movement
toward the large domain. As shown in Figure 6C, the M44
side chain is inserted into the hydrophobic pocket and
forms a contact with the P243 side chain. M47 cannot
form this interaction with the hydrophobic pocket without
Structure

forming improper bond angles in the context of these
rigid-body movements. The two residues (M44 and
P243) are simultaneously protected from solvent expo-sure
through the ‘‘pairwise’’ interactions. On the other
hand, M47 is exposed to solvent almost completely.
This makes predictions that could be tested in future foot-printing
experiments using high-resolution tandem MS.
In systematic mutational analysis experiments on yeast
actin, the double-mutation A204E/P243K abolishes poly-merization
(Joel et al., 2004). These mutations introduce
bulky and charged residues at the place where contacts
are formed in our model, likely blocking this conforma-tional
change. However, because these residues are also
involved in the intermolecular contacts of the F-actin
model, it is also possible that the mutations may hinder
F-actin filament assembly.
The closure of the large cleft in our model allows the
formation of a hydrogen-bond network between SD2
and SD4 (Figure 6D). In our model, seven new hydrogen
Figure 6. Magnified Views of Interface
between SD2 and SD4
(A) Template structure 1YAG.
(B) The model. The residues that experience
protection are colored blue (in SD2) or red
(SD4).
(C) The interaction between the SD4 hydropho-bic
pocket and D-loop M44 side chain. The
side chain of P243 is colored green while the
side chain of M44 is displayed as bubbles (sul-fur
is colored yellow).
(D) Newly formed hydrogen-bond network be-tween
SD2 and SD4 in the model. Residues are
silver in SD2 and brown in SD4. The possible
p-stacking is indicated by surface models on
residues Y69 and R183.
bonds can be formed between SD2 and SD4, as indicated
in the figure. Also, Y69 and R183 are in good locations for
p-stacking interactions (this interaction may not be critical
to form the structure, as the R183A/D184A mutant has no
significant change in phenotype). In a comparison of ASA
between 1YAG (rabbit sequence) and the model (Figure 7),
M47’s side-chain ASA increased by almost 50%. On the
other hand, M44 experiences almost complete burial of its
side chain. If we take into account the extremely dynamic
nature of the D loop, it is reasonable to assume that
both M44 and M47 are much more accessible than the
structure observed in 1YAG. Overall, the accessibility of
M44/47 seen in footprinting is in good agreement with
the model. The ASA changes in Y53 can be explained in
the same way. The D-loop structure is highly mobile, and
in the crystal structure the side chain of Y53 in 1YAG is
covered by the D loop and appears buried, but this does
not reflect the ensemble average experienced by the fluc-tuating
structure. In the model, although the ASA of the
Figure 7. Changes in Accessible Surface
Areas of Probe Residues
The red bars are ASAs of 1YAG (rabbit) and the
blue bars are differences between 1YAG and
the model. Negative values indicate less ex-posed
in the model.
Structure

Y53 side chain increases about 25A°
2, the D-loop structure
should be stiff through its formation of an interaction with
SD4. Thus, if the dynamic nature of structural changes in
SD2 is considered, this residue can experience a lower
reactivity consistent with the structural changes seen in
the model.
DISCUSSION
Possible Mechanism of Cleft Closure
Our model is based on rigid-body movements of domains
and subdomains observed in solved crystal structures.
We applied changes strictly to the backbone angles that
are involved in domain movements because we do not
have any data that indicate there is large-scale reorgani-zation
of structure other than domain movements. The
large-domain movement relative to SD1 narrowed the
large cleft including the nucleotide binding pocket as pro-posed
in previous reports of F-actin structure (Belmont
et al., 1999; Lorenz et al., 1993; Tirion et al., 1995). In the
crystal structures except 1RFQ-B, the metal ion is coordi-nated
only by water molecules and phosphate oxygen
atoms. The narrowed cleft results in a closer proximity of
side-chain oxygen atoms (D11:OD1, OD2, and Q137:OE1)
to metal ions that coordinate phosphate oxygen atoms. In
our model, the side chains of these residues form inner-sphere
coordination to Mg2+. AsMg2+ ions strongly prefer
to coordinate to oxygen atoms of side chains instead of
those of water molecules with a likely gain of free energy
(Dudev et al., 1999), it is reasonable to assume that ex-change
of inner-sphere coordinations between water mol-ecules
and oxygen atoms in D11 and Q137 occurs spon-taneously,
as thermal fluctuations bring side-chain oxygen
atoms close enough to the metal ions mediating the ex-change
reaction. This can then be a driving force for the
movement of the large domain toward SD1. Interestingly,
there is no strong interaction between SD1 and SD3—no
hydrogen-bond network nor hydrophobic interactions.
Thus, there is no discernable preference (or penalty) for
these subdomain motions compared to the structures ob-served
in the crystals. Once the large domain is locked in
the closed form by formation of inner-sphere coordination
between side-chain oxygen atoms and the metal ion, SD2
and SD4 are closer to each other and in the range where
a detailed hydrogen-bond network (and possibly p-stack-ing)
can form between them. This hydrogen-bond network
and p-stacking stabilize and lock a ‘‘super-closed’’ form
of the large cleft. The invariant residue Y69 appears to
be a key residue for the interaction between SD2 and
SD4. It possibly forms hydrogen bonds with the residues
in SD4 at the bottom of the large cleft and acts as a latch.
Mutations of residues forming interactions between SD2
and SD4 (Figure 6D) are reportedly lethal in yeast (Sheter-line
and Sparrow, 1994; Wertman et al., 1992) (R62, R206,
E207), although these residues located within the cleft are
unlikely to be involved in protein/protein interactions.
The contribution of the D loop to cleft closure is
documented in the literature (Khaitlina and Strzelecka-
Golaszewska, 2002). The cleavages in the D loop by
ECP32 protease (G42-V43) or subtilisin (M47-G48) greatly
stimulate the tryptic susceptibility of residues within SD2
(R62 and K68). The cleavage of the D loop apparently
makes those accessibilities of tryptic cleavage sites in-crease.
Thus, the D loop contributes to the stability of the
closed structure although it is not clear how much contri-bution
to the stability comes from the D loop. Our model
predicts the interaction between M44 and the SD4 hydro-phobic
pocket contributes to stabilization of the closed
structure of the large cleft. The observations above sup-port
our model, as the D-loop/SD4 interaction is a strong
candidate for the change in dynamics of SD2.
Catalytic Residue
As a consequence of the large-cleft closure by large-domain
movements relative to SD1, the geometry inside
the nucleotide binding pocket is altered from those seen in
typical crystal structures. The narrowed cleft brings D11,
Q137, and D154 side chains closer to the metal ion and
the b and g phosphates while maintaining the hydrogen-bond
network already in place.
D11 and Q137 are likely to be involved in the inner-sphere
metal-ion coordination. On the other hand, D154
seems to be too far from the metal ion to coordinate but
is located in a good position to form an inline attack of the
g phosphate. The double-mutant D154A/D157A is lethal in
yeast (Wertman et al., 1992); however, the S14C/D157A
double mutant does not show significant change in
ATPase activity (Schu¨ ler et al., 1999). Thus, it is very likely
that the D154A mutation is directly involved in lethality. As
suggested in the literature (Schu¨ ler, 2001), D154 appears
to be a prime candidate for the catalytic residue from both
previous reports and our model.
Actin Model Consistency with Nucleotide
and Metal-Ion Exchange Rates
The stability constants of the Mg2+- and Ca2+-ATP com-plexes
are similar (4.2 and 4.0, respectively) (Williams,
1970), and the exchange rate constants of the metal
ions and ATP in water are quite rapid (3.3 3 105 and 2.0 3
105 s1) (De La Cruz and Pollard, 1995; Pecoraro et al.,
1984). On the other hand, the exchange rates of Mg2+
and Ca2+ bound to ATP-G-actin are quite slow compared
with the rates observed in water and are different from
each other (association rates: 2.3 3 105 and 2 3 107
M1 s1; dissociation rates: 0.0015 and 0.014 s1 for
Mg2+- and Ca2+-ATP-G-actin, respectively, as summa-rized
in Table 3) (Gershman et al., 1991; Selden et al.,
1989; Sheterline and Sparrow, 1994). These data indicate
that the metal ions are significantly stabilized within the
cleft by limited diffusion or coordination bond formation
with protein side-chain oxygen atoms. The difference in
association/dissociation with ATP between Mg2+ and
Ca2+ is not significant in water but is an order of magnitude
different in the case of actin-nucleotide-bound forms. Our
model predicts that one factor explaining the difference in
stability is coordination of Mg2+ with oxygen atoms of the
protein. The other factor is the difference in accessibility of
metal ions (steric effect). The exchange rates of ATP for
Structure

Table 3. Exchange Rate Constants of Nucleotides/Metal Ions for Different Species of Actin
Rate Constant Mg2+ Ca2+ Species
Metal ion Exchange rate 3.3 3 105 s1 2.0 3 105 s1 ATP/metal ion
Metal ion Association rate 2.3 3 105 M1 s1 2.0 3 107 M1 s1 Metal ion/ATP + actin
Metal ion Dissociation rate 0.0015 s1 0.014 s1 Metal ion/ATP/actin
ATP Exchange ratea 5.0 3 104 s1b 1.5 3 102 s1c Metal ion/ATP/actin
Metal ion Affinity (Kd) 10 nM 2 nM ATP-G-actin
a The exchange rate is highly dependent on metal-ion concentrations.
bMg2+ concentration of 1 mM.
c Ca2+ concentration of 0.1 mM.
both Mg2+- and Ca2+-G-actin are even more different (5 3
104 and 1.5 3 102, respectively) (Sheterline and Spar-row,
1994). As ATP exchange rates are tightly connected
to free metal concentrations, the metal-ion exchange
rate would be a limiting factor for nucleotide exchange
(Kinosian et al., 1993). Even if we attribute all of the metal
exchange rate difference to coordination states, still there
is a difference in ATP exchange rates. Thus, this ATP ex-change
rate difference may be related to the steric effect
of the cleft closure. These observed exchange rate differ-ences
between Mg2+- and Ca2+-G-actin are quite consis-tent
with our model (closed cleft and direct coordination of
Mg2+ to the side-chain oxygen atoms).
Implications for F-Actin Models
The Holmes model of F-actin (Holmes et al., 1990; Lorenz
et al., 1993; Tirion et al., 1995) has been widely accepted
and is largely consistent with X-ray fiber diffraction (Bel-mont
et al., 1999; Holmes et al., 1990; Lorenz et al.,
1993; Oda et al., 2001; Tirion et al., 1995; Wu and Ma,
2004), cryoelectron microscopy (Schmid et al., 2004),
and atomic force microscopy data (Shao et al., 2000; Shi
et al., 2001). We have examined the F-actin structure by
side-chain solvent accessibility utilizing hydroxyl-radical
footprinting (Guan et al., 2005). Although our data were
mainly consistent with the F-actin model, there are some
disagreements with the Holmes model. The most appar-ent
difference is the status of the large cleft. The protection
sites in SD4 (residues 200–202 and 243) and the D loop
(residues 40, 44, and 47) form contacts in the intermolec-ular
interfaces; there is no disagreement with footprinting
data for these residues. However, the hydroxyl-radical re-activity
within the large cleft is significantly lower in F-actin
compared with Ca2+-G-actin. A closed cleft is also ob-served
in a reconstructed model from cryo-EM data
(Schmid et al., 2004), although the degree of closure
seems not to be enough to explain the almost 90% reduc-tion
in hydroxyl-radical reactivity compared to the open
form of Ca2+-G-actin.
The ADP and ADP-BeF3
forms of fiber diffraction data
show differences in the status of the large cleft (Belmont
et al., 1999); the latter form indicates a more closed cleft.
The replacement of ADP with ADP-BeF3
also diminishes
susceptibility of F-actin to tryptic cleavage within SD2
(Muhlrad et al., 1994). As ADP-BeF3
mimics ATP or
ADP-Pi (Muhlrad et al., 1994), this form represents ATP
or ADP-Pi-F-actin before it releasesPi. These observations
indicate that in the F-actin filament, hydrolysis of ATP to
ADPand subsequent release of Pi correlate with large-cleft
opening. On the other hand, when Mg-ATP-G-actin is in-corporated
into the filament, we suggest it is in the highly
closed form seen here; this Mg-ATP form of actin has a
lower critical concentration than Ca2+-ATP-G-actin. Catal-ysis
mediated by the Mg2+ ion may drive conformational
changes among the coordination bonds between side
chains and the metal ion, leading to rearrangements of
subdomains 2 and 4 and the generation of a more open
cleft within the ADP-F-actin structure. This would provide
a more flexible and dynamic SD2, which is thought to be
a key factor in attracting binding proteins and driving fila-ment
dynamics (Schmid et al., 2004).
Conclusion
Understanding monomeric G-actin structure is important,
as it is relevant to many protein/protein interactions in
which actin participates. Hydroxyl-radical footprinting is
a powerful technique to probe surface accessibility of
macromolecules. The technique is a local measure as it
reports side-chain accessibility. Such information pro-vides
strict restraints for estimating changes in structure
and in defining domain interactions. In conjunction with
a computational approach, a structural model of the Mg-
ATP-actin monomer was built that is consistent with ex-perimental
data. This model provides novel insights into
the conformational rearrangements driving actin filament
dynamics and provides a unique structural insight into ac-tin
monomer structure. This combined computational and
experimental approach is particularly useful for modeling
the structural changes of proteins when we do not have
access to crystal or solution structures for conformations
of functional interest, but where we may have suitable
starting templates for modeling the structure. Further
developments in computational modeling will ease the
integration of the experimental into the computational
approach as a filtering/validation tool.
EXPERIMENTAL PROCEDURES
Rigid-Body Movement Analyses
Tcl scripts for molecular viewer VMD (Humphrey et al., 1996) were de-veloped
for the analysis and visualization of rigid-body rotational
Structure

movements of domains/subdomains. The scripts accept the range of
the target residues in two structures for rigid-body movement and then
calculate vectors of atom movements and averaged normal for these
vectors; they then search for the atoms within the reference structure
that are nearest to the center of motion. The algorithm is based on the
assumption that there are pivot points in bonds within the backbone
allowing rigid-body rotational movements (the movements should be
explained by rotations of some bond angles). The algorithm is based
on the following step-by-step calculations.
Define structures A and B, where A is the reference structure and B
is the target structure for movement calculation. The total number of
atoms(Ca orbackbone atoms) isn within the target range.The totalnum-ber
of the atoms in reference structure A (Ca or backbone atoms) is N.
v !
Atoms in structure A: ai (i = 1 to n), coordinates Pai.
Atoms in structure B: bi (i = 1 to n), coordinates Pbi.
Atoms for the center C: cj (j = 1 to N), coordinates Pcj.
The midpoint of ai and bi: mi (i = 1 to n), coordinates Pmi.
The vector between points ai and bi: i = Pbi Pai .
The vector between points mi and cj: u !
ij =Pmi Pcj .
The angle formed by the two vectors v !
i and u !
ij :
qij =

acos

ð v !
i u !
jiÞ
jv !

i j
u !
ji

:
The averaged angle for the point Pcj:
q
j =
1
n
Xn
i =1

qij
p
2

ð j =1toNÞ:
The SD of the angles for atom Pcj:
SDj =
ffiPffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi n
i =1ððqij p
2

q
j
2
n 1
s
ð j =1toNÞ
The averaged angle between the real center of movement cr and the
2; thus, q
two atoms in structures A and B should be p
j is 0. We need to
find the atom that gives the averaged angle closest to 0. In order to
avoid the cases where q
j happens to be close to 0 but angles vary
as qij can be positive or negative, the score is calculated by the follow-ing
formula:
Sj =q
j 3SDj :
Then, the script calculates the average positions of the top scored
five atoms. This point is defined as gc.
Now, we need to calculate the normal vector for the rotational move-ments.
The normal vector for the rotational movements should be ap-proximately
perpendicular to all vectors from the center of movement
to the midpoint of atoms in the two structures A and B. The normal vec-tor
is also perpendicular to the vector vi between ai and bi. In order to
minimize the contribution from small vectors that tend to have larger
deviation from ideal ones (the vectors are extended by unitization
along with ‘‘error’’), ‘‘normalized’’ normal vectors are weighed by their
norms. After the average normal vector is calculated, it is renormalized
to the unit vector.
The vector between points mi and gc: w !
i =Pmi gc.
The normal vector of rotation for points ai and bi:
n !
i =
v !
i 3 w !
i
jv !
i jjw !
i j
(crossproduct).
Averaged (weighed) vector:
n !
av =
1
n
Xn
i =1
i jjw !
i j,n !
ðj v !
iÞ:
Normal vector (unit vector):
n !
unit =
n !
av
jn !
:
av j
The resultant averaged normal vector and center of rotation is
visualized along with vectors vi within VMD by Tcl commands in the
script.
Accessible Surface Area Calculations
The accessible surface areas of side chains are calculated using the
GETAREA 1.1 (Fraczkiewicz and Braun, 1998) web server with 1.4 A °
probe radius (http://www.scsb.utmb.edu/cgi-bin/get_a_form.tcl) with
additional atom type library and residue type entries. In our previous
report, we used rabbit skeletal muscle actin. As ASA greatly depends
on the side chain, the sequence of rabbit skeletal muscle actin was
mapped on 1YAG using MODELLER 7v7(Sali and Blundell, 1993).
This structure is designated as 1YAG (rabbit sequence) in the following
sections.
Computational Modeling
Model construction started from a ‘‘template’’ structure of 1YAG (rab-bit
sequence). The rotational movement toward the small domain was
applied to the large domain (residues 138–332) at the pivot residue 138
with the normal vector calculated in 1HLU (Chik et al., 1996). The rota-tional
movement toward SD2 was applied as a combination of several
movements calculated based on the analyses of several crystal struc-tures.
The rotations were applied over four residues from specifically
selected pivot points (residues 34–37 and 66–69) in order to prevent
too much rotation to single bonds that can lead to improper bond an-gles.
All rotational transformations of coordinates were performed us-ing
Tcl scripts within VMD. Modeled structures were generated with
combinations of different angles and normal vectors. ASAs of side
chains were calculated for these structures in order to assess the pre-liminary
models. The best model in the series of models by rigid-body
movements was chosen according to the consistency with expected
changes in ASAs of hydroxyl-radical probes.
The model was then subjected to manual inspections/manipulations
of side chains in the Swiss-PdbViewer (Guex and Peitsch, 1997) in or-der
to better satisfy the hydroxyl-radical data. The rotamer of side
chains was selected to maximize hydrogen-bond formations within
the large cleft. As the large cleft between SD2 and SD4 was closed
by moving the large domain relative to SD1, the nucleotide binding
pocket was also affected and became significantly narrower than in
the template structure 1YAG. The coordinates of the phosphates of
ATP were manually adjusted within VMD using Tcl scripts by rotating
the phosphate-oxygen bond and fit into the narrower cleft interactively
in order to avoid steric clashes. The D loop was constructed manually
in the Swiss-PdbViewer by rotating the phi and psi angles of the tem-plate
structure. This process was guided mainly to allow M44 to reach
the hydrophobic pocket in SD4 while at the same time avoiding im-proper
angles. At this stage, the model structure was examined by
PROCHECK (Laskowski et al., 1993) and Verify3D (Bowie et al., 1991)
for model validity (Figure S1). The model coordinates were then passed
to MODELLER 7v7 (Sali and Blundell, 1993) as a template for adjusting
and correcting the stereochemical parameters. MODELLER was also
used to adjust the geometry of the residues and metal ion in the nucle-otide
binding pocket with manually assigned restraints according to the
1RFQ-B (chain B) structure (Reutzel et al., 2004). After further manual
inspections/adjustments in VMD and energy minimization within the
Swiss-PdbViewer for certain side chains, the final structural model
was subjected to PROCHECK and Verify3D for the assessment of
structure validity.
Visualization of Structures
The structures were visualized using POV-Ray ray-tracer (Version 3.6)
with output from VMD (Version 1.8.3) and manual editing of scene files.
Some presentations were directly defined by POV-Ray scene. All op-erations
were done on a Power Macintosh G5 with Mac OS X except
for energy minimization within the Swiss-PdbViewer that was per-formed
on a Windows 2000 workstation (OS X version is an a version
and energy minimization is only partially implemented).
Structure

Supplemental Data
Supplemental Data include two figures and Supplemental Results
and are available at http://www.structure.org/cgi/content/full/15/1/
39/DC1/.
ACKNOWLEDGMENTS
This work is supported in part by the Biomedical Technology Centers
Program of the National Institutes for Biomedical Imaging and Bioen-gineering
(P41-EB-01979).
Received: July 26, 2006
Revised: November 6, 2006
Accepted: November 18, 2006
Published: January 16, 2007
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