Biopolymer s
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Transcript of Biopolymer s
Relating Nucleotide-Dependent Conformational Changes in Free TubulinDimers to Tubulin Assembly
Kathiresan Natarajan, Jagan Mohan, Sanjib SenapatiDepartment of Biotechnology, Indian Institute of Technology Madras, Chennai 600036, India
Received 12 June 2012; accepted 29 August 2012
Published online 10 September 2012 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/bip.22153
This article was originally published online as an accepted
preprint. The ‘‘Published Online’’ date corresponds to the
preprint version. You can request a copy of the preprint by
emailing the Biopolymers editorial office at biopolymers@wiley.
com
INTRODUCTION
Microtubules (MTs) are cytoskeletal polymers that
play key roles in diverse array of cellular func-
tions, such as mitosis and meiosis, motility,
maintenance of cell shape, and intracellular
transport of organelles. The a,b-tubulin dimer is
the basic structural unit of MTs. Essential to the microtubule
stability and function, the dimer contains two nonidentical
guanine-nucleotide binding sites—a nonexchangeable (N)
site at a-tubulin that always binds a GTP molecule, and
an exchangeable (E) site at b-tubulin that can bind either a
GTP or GDP.1,2 The bound nucleotide at E site stays
in equilibrium with free nucleotides in cytosol, where the
GDP $ GTP exchange at E site can take place freely.3,4
In vitro, MT assembly from purified tubulin solutions
involves polymer nucleation and elongation to reach a steady
state, which is characterized by a constant ratio of polymeric
tubulin to free tubulin dimers. According to the existing
model, nucleation is the crucial initial phase where several
GTP-tubulin dimers assemble to form oligomers that subse-
quently combine to form 2D sheets and microtubules.5,6
Thus, a detailed knowledge of the structural differences
between free GTP- and GDP-tubulin dimers in solution
has direct relevance in understanding the MT assembly and
function.
Relating Nucleotide-Dependent Conformational Changes in Free TubulinDimers to Tubulin Assembly
Additional Supporting Information may be found in the online version of this
article.Correspondence to: Sanjib Senapati; e-mail: [email protected]
ABSTRACT:
The complex dynamic behavior of microtubules (MTs) is
believed to be primarily due to the ab-tubulin dimer
architecture and its intrinsic GTPase activity. Hence, a
detailed knowledge of the conformational variations of
isolated a-GTP-b-GTP- and a-GTP-b-GDP-tubulin
dimers in solution and their implications to interdimer
interactions and stability is directly relevant to
understand the MT dynamics. An attempt has been made
here by combining molecular dynamics (MD)
simulations and protein–protein docking studies that
unravels key structural features of tubulin dimer in
different nucleotide states and correlates their association
to tubulin assembly. Results from simulations suggest that
tubulin dimers and oligomers attain curved
conformations in both GTP and GDP states. Results also
indicate that the tubulin C-terminal domain and the
nucleotide state are closely linked. Protein–protein
docking in combination with MD simulations suggest
that the GTP-tubulin dimers engage in relatively stronger
interdimer interactions even though the interdimer
interfaces are bent in both GTP and GDP tubulin
complexes, providing valuable insights on in vitro finding
that GTP-tubulin is a better assembly candidate than
GDP-tubulin during the MT nucleation and elongation
processes. # 2012 Wiley Periodicals, Inc. Biopolymers 99:
282–291, 2013.
Keywords: tubulin dimer; nucleotide state; tubulin
assembly; molecular dynamics simulation; protein–
protein docking
Contract grant sponsor: Council of Scientific and Industrial Research (CSIR)
VVC 2012 Wiley Periodicals, Inc.
282 Biopolymers Volume 99 / Number 5
Although crystal structures of a,b-tubulin dimer bound
to different antimitotic agents are available,1,2,7–13 till date
there is no report of experimentally determined structures of
isolated GTP- and GDP-tubulin dimers. Such structures will
be of immense interest since predominantly the isolated
tubulin dimers regulate the microtubule function, as dis-
cussed above. The crystal structures of tubulin show three
functionally distinct domains in each monomer (Figure 1).
An N-terminal domain (N-domain) that constitutes the nu-
cleotide-binding region, an intermediate domain (I-domain)
that contains the taxol binding site, and a C-terminal domain
(C-domain) that has been implicated in the binding of sev-
eral motor proteins. The N-terminal domain comprises of
alternating parallel b-strands (S1–S6) and a-helices (H1–
H6). The loops, T1–T6 that join each strand and helix along
with the core helix H7, form the nucleotide binding pocket.
The intermediate domain formed by three helices (H8 –
H10) and a mixed b sheet (S7 – S10), involves in lateral and
longitudinal contacts with neighboring dimers. The loop
between S7 and H9, commonly known as M loop, comprises
part of the taxol-binding site in b-tubulin. The loop between
helices H7 and H8 (loop T7) and H8 itself are seen to inter-
act with the nucleotide of next subunit along the protofila-
ment. The C-terminal domain is formed by two antiparallel
helices, H11 and H12 that fold across the other two domains.
The small helix, H11’ connecting these two helices is also im-
portant for the interaction with the next monomer along the
protofilament.
Computational methods have proven to be a valuable tool
for biological research. Recent computational studies have
shed light on various properties of tubulins and MTs, which
were otherwise difficult to explore experimentally.14–30 The
role of nucleotides on structural rearrangements of tubulin
dimers has recently been investigated by Gebremichael et
al.21 and Bennett et al.22 These simulation studies have indi-
cated intrinsic bending in tubulin dimers. Bennet et al. also
have noted that the dimer flexibility plays an important role
in tubulin assembly. Simulations of model protofilaments
have helped to understand the structural features of tubulin
oligomers present in solution,23 binding of drugs24,25 and
antimitotic peptides,26,27 and the elastic properties of tubu-
lins in assembly.28–30
In this study, we use all-atom MD simulations and pro-
tein–protein docking to explore the binding characteristics of
tubulin dimers to produce tubulin oligomers, in different
nucleotide states. Specifically, we aim to elucidate the struc-
tural variations that make GTP-tubulin a better assembly
candidate than GDP-tubulin during MT nucleation and
elongation processes. Results suggest that GTP fine-tunes the
functional loop regions at interdimer interface to achieve
better binding with the next GTP-tubulin. Results also imply
that the tubulin C-terminal domain and the nucleotide state
are closely linked. Moreover, GTP-tubulins are found to
engage in relatively stronger interdimer interactions even
though the interdimer interfaces are bent in both GTP and
GDP tubulin oligomers.
RESULTS AND DISCUSSIONSTo begin with, we have performed two all-atom MD simula-
tions of ab-tubulin dimer liganded to GTP at N site and
GDP or GTP at E site. The simulations were initiated from
the straight crystal structure of tubulin dimer bound to taxol
(PDB ID: 1JFF1). Taxol was removed from the crystal struc-
ture, and the resultant conformation was equilibrated in
explicit water via 20 ns MD run, to obtain a model structure
for a-GTP-b-GDP-tubulin dimer. This equilibrated structure
was further engineered to generate the starting structure for
the second simulation, where we modified the GDP at E site
to GTP (by this means we wanted the GDP-to-GTP-exchange
to take place without the influence of taxol, as presumably
happens in cytosol). The resultant structure was then equili-
FIGURE 1 Taxol-bound straight crystal structure of ab-tubulin
heterodimer (PDB ID: 1JFF). For clarity, only the functionally rele-
vant loops T2, T3, T4, T5, H1-S2 from N-domain (red); loops M,
H6-H7, T7, helix H8 from I-domain (purple); and helices H11, H12
from C-domain (blue) are highlighted.
Nucleotide-Dependent Changes in Free Dimers to Tubulin Assembly 283
Biopolymers
brated via 20 ns MD run to obtain a model structure for a-
GTP-b-GTP-tubulin dimer. Both the model structures were
further simulated to generate 250 ns MD data, upon which
all analyses were performed. The simulation details are pre-
sented under ‘‘Materials and Methods’’ section.
Conformational Changes in GTP- and GDP-Tubulin
Dimers
The b-subunit is found to undergo relatively larger confor-
mational changes than the a-subunit in both GTP-tubulin
and GDP-tubulin simulations (Supporting Information Fig-
ure S1). This is not surprising, considering the fact that both
taxol binding and GTP-hydrolysis take place within the b-
subunit. Moreover, a comparison of the root mean squared
displacements (RMSD) of the residues of b-subunits implies
that the GTP-tubulin residues deviate slightly more than the
GDP-tubulin residues from the starting crystal conformation
(Supporting Information Figure S2). A detailed comparison
of the model structures with the crystal conformation is pre-
sented in Figure 2. The figure is generated by a stereo super-
position of the crystal structure and the final 100 ns time-
averaged structures from simulations, according to Ca atoms
of the b-subunits. In a 3D representation of the structural
elements, the figure highlights the most notable variances
among the three states of tubulin dimer. The sites of maxi-
mum variances include the nucleotide binding loops T1, T2,
T3, T5, and some of the allosteric loops, M loop, H6-H7
loop, H1-S2 loop in b-monomer. The T1, T2 loops close
down over the nucleotide in GTP-tubulin under the influ-
ence of additional c-phosphate. The T3 and T5 loops, which
are known to be involved in longitudinal contacts, show
larger deviation in GTP-tubulin dimer. The M loop that
comprises part of the taxol binding site, exhibits a great deal
of variation. Although it is found to protrude outward later-
ally in GTP-tubulin dimer as similar to the taxol-bound
tubulin structure (though in larger extent in the former), it
protrudes inward in GDP-tubulin dimer. The H1-S2 loop
that resides opposite to M loop also demonstrates a greater
outward movement, when the protein b-subunit is bound to
GTP or taxol. The H6-H7 loop, which is known to play a key
role in longitudinal interactions along the protofilaments,
shows greater extendibility in GTP-tubulin dimer than in the
other two systems. These changes are shown in Figure 2. To
visualize the differential movements of these loops more
clearly, we have plotted their distances from the core helix
H7 as a function of time in GTP and GDP states. As Figure 3
indicates, these loops are more flexible and tend to extend
farther in GTP state. The greater tendency of some of these
loop regions for outward longitudinal movements in GTP-
tubulin might suggest that GTP binding favors interdimer
interactions and hence the protofilament elongation. It is
generally believed that largely extended protein residues are
more free to move and hence can readily approach to the
neighboring subunits for stronger interactions. It is also
reported that GTP binding favors protofilament elonga-
tion.31 The movements of the allosteric loops due to nucleo-
tide exchange are not simply stochastic, as shown by recent
mutational and protein–drug interaction studies where
strong correlations between tubulin N- and I-domains were
noted.24,32
When this work was in progress, the crystal structures of
GTP- and GDP-tubulins in complex with a stathmin-like do-
main were reported.31 In accordance with the crystal struc-
tures, our simulation results show a great deal of difference
between GTP- and GDP-tubulins in the T3 and T5 loop
region. Both the loops, in GTP-tubulin, show greater propen-
sity to involve in longitudinal interactions by extending out-
ward longitudinally (Figures 2 and 4a). The direct interaction
FIGURE 2 Time-averaged structures of the model GTP- and
GDP-tubulin dimers, superposed on the taxol-bound tubulin crystal
structure, based on the Ca atoms of b-subunits. Secondary struc-
tural elements which underwent the most significant conforma-
tional changes are highlighted. Color scheme—orange: GTP-tubu-
lin, green: GDP-tubulin, blue: crystal structure.
284 Natarajan, Mohan, and Senapati
Biopolymers
of GTP c-phosphate with the T3 loop helps the b:Asn101
side chain swinging out, which in turn modulates the posi-
tion of b:Thr180 in T5 loop. As a result, the H-bond between
b:Asn101 and b:Thr180 breaks and the T5 loop flips out in
GTP state, as similar to the crystal structure of GTP-tubulin.
This makes b:Asn101, which is the most conserved residue
across the species and known to make longitudinal contact,
more available for interactions. The T5 loop residue
b:Asp179 is also seen to become more exposed in GTP-tubu-
lin, similar to the crystal structure. These changes are shown
in Figure 4a and Supporting Information Movies S1 and S2.
Our simulation data could also capture both ‘‘in’’ and ‘‘out’’
conformations of T5 loop in GDP-tubulin as similar to the
crystal structure, even though the flip-out movement was
short lived (Figure 3b).
It is noteworthy that the core elements of the protein also
have undergone changes under the influence of the nucleo-
tide. This is shown in Figure 4b, where the superposition of
N- and C-terminal domains of the b-subunit demonstrates
variations in the intermediate domain. This domain is found
FIGURE 3 Differential movements of functionally relevant loops in GTP and GDP states. A dis-
tance metric is used, in which the Ca distance of the central residue of a loop is measured from the
centre of core helix H7, over time. The regions of interest include: (a) T3 loop, (b) T5 loop, (c) H6-
H7 loop, (d) M loop, and (e) H1-S2 loop. Color scheme—Red: GTP-tubulin; green: GDP-tubulin.
All the loop regions appear to be more flexible and to extend farther in GTP state. In (b), the T5
loop of GDP-tubulin attains ‘‘out’’ conformation similar to the GTP state during 0–30 ns and 100–
130 ns of simulation time. Otherwise, this loop attains ‘‘in’’ conformation in GDP state.
Nucleotide-Dependent Changes in Free Dimers to Tubulin Assembly 285
Biopolymers
to rotate slightly toward N-domain in GTP state but rotates
in opposite direction in the GDP state. A very similar rota-
tion in the intermediate domain was noted, while comparing
the classical curved (PDB ID: 1SAO) to straight tubulin
structures (PDB ID: 1JFF) by tubulin N- and C-terminal
superposition. The observed change in the intermediate
domain is also consistent with the proposal of Amos and
Lowe that ‘‘hydrolysis (GTP to GDP state) could vary the rel-
ative orientation of the N terminal domain and intermediate
domain.’’33
Nucleotide-Dependent Intradimer Bending
The controversy over straight versus curved conformation of
ab-tubulin dimer remains unresolved over the decades.
Although the allosteric model postulates that GTP-tubulin
dimer is straighter and prestructured in solution for assembly
onto the microtubule wall,34–37 the lattice model proposes
that the tubulin dimers adopt the curved conformation in
solution irrespective of the nucleotide state.38–43 In this
work, following the prescription of Nogales and Wang,36 we
have estimated the intradimer bending in tubulin by com-
puting the angle subtended by the axis of two monomers
within the dimer. The bending deformation is quantified by
computing the instantaneous angle assumed by two vectors
A and C, drawn on the molecular frames. The origin of the
coordinate system was chosen to be the centre of mass
(COM) of the overlapping region of a- and b-monomers,
primarily comprised of a:H6-H7, a:H11-H12, a:T5, a:T3,
a:T2, b:H10, b:H10-S9, b:S9, b:H8, and b:T7. The vector
from the origin to the COM of a-subunit (excluding the resi-
dues considered in overlapping region) was termed as A, and
the one from the origin to the COM of b-subunit (excluding
the residues in overlapping region) was termed as C. The vec-
tors are set to point to the COM of the subunits. The angle
subtended by the axes of two subunits was then computed as:
H ¼ 180o � Cos�1ðA:CÞ=jAjjCj:
The computed angular distributions of the model dimers,
sampled over the final 100 ns production phase, are pre-
sented in Figure 5. The result from the control simulation of
taxol-bound GDP-tubulin crystal structure is also included
for comparison. The profiles show a similar shape with the
distribution nearly Gaussian, peaking at an angle of 208 for
GDP-tubulin and 178 for GTP-tubulin. The taxol-bound
structure is seen to distribute around 148, in comparison to
its value of 118 in the lattice-constrained conformation in the
crystal structure. The angle values thus indicate that tubulin
relaxes from straight crystal conformation to more curved
conformations in isolated states and both the GTP and GDP
tubulin dimers are naturally bent. This finding is consistent
with the recent crystal structures, where both GTP- and
GDP-tubulins, bound to RB3-SLD protein, were found to
attain curved conformations.31 The advantage of the present
simulation data is that it covers a range of distribution of the
intradimer angle, out of which the curved conformation
stands out to be the most probable state. The time-averaged
FIGURE 4 (a) Close-up view of the nucleotide binding region.
Loops T3, T5 from both GTP- (orange) and GDP-tubulins (green)
are highlighted. Also shown is the nucleotide in GTP state. The
presence of an Mg21 ion along with its coordinating water was
observed around the GTP in GTP-tubulin. (b) Change in intermedi-
ate domain (orange: GTP-tubulin, green: GDP-tubulin) is high-
lighted by superposing the N- and C-domains (light green).
286 Natarajan, Mohan, and Senapati
Biopolymers
structures of the model GTP- and GDP-tubulins are com-
pared with the curved crystal structures of tubulin dimer
from the work of Nawrotek et al.31 and are shown in Sup-
porting Information Figure S3. The RMSD values for the
GTP- and GDP-states are only 1.82 A and 1.64 A, suggesting
that tubulin dimer is indeed bent in both states. The direc-
tion of bending is outward, almost tangential to the MT wall.
Interestingly, the C-domain in GTP-tubulin dimer slightly
misaligns from the crystal structure. We will discuss it in a
subsequent section.
Nucleotide State and Tubulin Assembly
Recent electron microscopy studies due to Mozziconacci et
al.44 have suggested that the tubulin dimers can exist as short
oligomers in solution. Moreover, tubulin oligomers form the
nuclei for MT nucleation process.5,6 These reports encour-
aged us to construct the tubulin dimer–dimer complexes
from the model tubulin structures and investigate their bind-
ing characteristics in GTP- and GDP-states. The dimer–
dimer complexes were initially built from the time-averaged
structures, using the protein–protein docking program
HADDOCK.45 The lowest energy complex with correct lon-
gitudinal pitch from each category was subjected to 50 ns
MD simulation for structure refinement. Figure 6 depicts the
time-averaged conformations of the complexes, obtained
from the final 20 ns simulation data. We found that the con-
formations of our model protofilaments resemble the RB3-
SLD crystal structure very closely, with a few additional lon-
gitudinal contacts. The calculated interdimer contact angle in
model GDP-protofilament is � 128, which is very close to
the value of 12.68 in the crystal structure. The GTP-protofila-
ment is also found to be bent with the interdimer angle is
� 88. The range of angles could be attributed to the plasticity
at the interdimer interface. It is also possible that all contacts
could not be formed in the docking experiment or during
the subsequent simulation time in GTP state.
It is worth mentioning here that, in the HADDOCK out-
put, even though the cluster-1 (i.e., energetically ranked 1) in
GTP-tubulin docking contained the lowest energy complexes
with correct pitch, in case of GDP-tubulin only cluster-3 and
cluster-4 (i.e., energetically ranked 3 and 4) contained the
conformations with correct longitudinal pitch. These cor-
rect-pitch-conformations represent 63% and 59% of the total
docked complexes in GTP- and GDP-state, with average
intermolecular energy 2678.85 6 30.74 and 2303.98 6
89.12 kcal/mol, respectively. For GDP-tubulin docking, the
lower energy conformations in cluster-1 and cluster-2 had
average intermolecular energies 2481.82 6 69.88 and
2366.81 6 66.47, respectively. In these dimer–dimer com-
plexes, however, the top dimer rotated substantially about
the vertical axis, which presumably could be an artifact of
defining the contact area through the choice of active
residues in HADDOCK. Hence, we warn the readers that
the output of HADDOCK is very sensitive to the choices of
FIGURE 6 Time-averaged structures of the dimer–dimer com-
plexes of (a) GTP-tubulins and (b) GDP-tubulins along longitudinal
direction. Secondary structural elements, primarily involved in lon-
gitudinal interactions include—loops H6-H7, H11-H12, T2, T3, T5
from b-tubulin (red) and loops H10-S9, T7, helices H8, H10 from
a’-tubulin (yellow). Also shown are the bound nucleotides, GTP
(magenta) and GDP (orange).
FIGURE 5 The distribution of intradimer bending angles in
model GTP- (solid line) and GDP-dimers (dashed line). The distri-
bution of taxol-bound structure is also included (dotted line).
Nucleotide-Dependent Changes in Free Dimers to Tubulin Assembly 287
Biopolymers
ambiguous intermolecular restraints (AIRs), and recommend
a detailed inspection of the output, plus a set of explicit sol-
vent MD simulations on the docking structures.
Figure 7 demonstrates the interdimer interface contacts
present in GTP- and GDP-tubulin complexes. The contact
map is obtained by calculating the areas of residue–residue
contacts at the dimer–dimer interface.46 As the figure indi-
cates, the total number of contacts in the two complexes is
nearly similar. However, a relatively stronger interdimer
binding in GTP state stems from the larger residue–residue
contact areas, as shown in Table S1. Table S1 lists the major
contacts present at the dimer–dimer interfaces, along with
the residue–residue contact areas. The total contact area at
the GTP and GDP dimer–dimer interface was found to be
2290 A2 and 1738 A2, respectively. The computed interface
contact areas are very similar to that in the crystal structure
(2550 A2; Ref. 31) and previously modeled protofilaments
(2000 A2; Ref. 23). The larger strength of GTP-tubulin longi-
tudinal contacts agrees well with their larger bending stiffness
in protofilaments, as noted by Grafmuller and Voth.23 This
finding is also consistent with in vitro studies, which impli-
cate GTP-tubulin to be a better assembly candidate than
GDP-tubulin during MT nucleation and elongation proc-
esses. Moreover, a closer look at Figure 7 indicates the direct
involvement of T5 and H6-H7 region from b-subunit of
lower tubulin dimer with helix H10 from a-subunit of upper
tubulin dimer in longitudinal assembly (the number of con-
tacts is, however, more in GTP state). This finding is also in
consistent with the proposal of Nawrotek et al. that GTP
binding promotes protofilament elongation, through the
‘‘out’’ conformation of T5 loop.31
Comparing the results to the previous MD studies of
tubulin dimers by Gebremichael et al. and Bennett et al. and
of tubulin oligomers by Grafmuller and Voth, it became evi-
dent that the intradimer bending angles found in this study
are larger than Gebremichael et al. and Bennett et al., but
comparable to the study of Grafmuller and Voth. This can
mainly be due to the fact that, while the former studies were
carried out for 20 ns or less, the later and this study are per-
formed for more than 100 ns. As noted by Grafmuller and
Voth and also in this study that, larger changes in bending
angle and direction often takes place on timescales of 50 ns
and longer. In this study, the direction of bending is seen to
be very similar to the direction described by Bennett et al.
and Grafmuller and Voth. The interdimer bending is also
comparable to the work of Grafmuller and Voth (88–128 in
this work versus 58–108 in Grafmuller and Voth).
Nucleotide States and Carboxy Terminal Domain
The carboxy terminal domain is an interesting but not thor-
oughly understood region of ab-tubulin dimer. Various b-
tubulin isoforms are known to vary sequence in this region.47
Further, this region serves as a binding site for microtubule
motors, associated proteins, and cationic molecules.48,49 Lit-
erature has suggested the possibility that the C-terminal do-
main and the nucleotide state of b-tubulin could be linked,
with the nucleotide state affecting the interaction capability
of the C-terminal region.50–52 Here, we attempt to explore
such a link by investigating the correlation between N- and
C-terminal domains in GTP and GDP states. This was done
FIGURE 7 Interdimer interface contacts in longitudinal dimer–
dimer complexes—(a) GTP-tubulin complex (b) GDP-tubulin com-
plex. Blue squares indicate the contact between pair of residues.
288 Natarajan, Mohan, and Senapati
Biopolymers
by examining the stability of the salt-bridge present at the
interface of N- and C-domain and also by measuring the
angle constituted by these two domains. The time evolution
of the salt-bridge, in Figure 8a, shows that the distance
between the bridging residues b:Lys105 in helix H3’ (N-do-
main) and b:Glu411 in helix H11’ (C-domain) is always sig-
nificantly smaller in GTP-dimer. This implies that the dislo-
cation of C-domain of GDP-dimer from straight orientation
in taxol-bound structure is more frequent. The distribution
of angles (Figure 8b), constituted by the vectors passing
through the interfacial helices H3’ and H11 also shows a dis-
tinct difference, where GTP-tubulin exhibits a bimodal distri-
bution. Although majority of tubulin conformations in GDP
state possess a C-domain bent by an angle of about 678,more than 50% of tubulin conformations in GTP state attain
a less-curved conformation with an angle of about 508. This
might suggest that GTP state prevents the outward curving
of C-domain in a greater extent, which thereby can engage in
more favorable interdimer association. The corresponding
angle found to be 65.38 in 1JFF and 78.68 and 79.38 in RB3-
SLD bound GTP- and GDP-tubulin crystal structures. The
similar and little larger angle values in GTP- and GDP-tubu-
lin crystals could be due to the bound RB3-SLD, which wraps
around the C-domain of tubulin.
SUMMARYAND CONCLUSIONSWe present a comprehensive study of tubulin dimers and
oligomers in different nucleotide states to examine the
intrinsic conformational changes in isolated dimers and
understand how these changes modulate the tubulin assem-
bly. Data show that functionally relevant loops, T3, T5, H6-
H7 exhibit greater tendency for outward longitudinal move-
ments in GTP-tubulin, suggesting that GTP binding can
facilitate interdimer interactions and protofilament elonga-
tion. In consistent with recent crystal structures, both GTP-
and GDP-bound tubulin dimers and oligomers are found to
have curved conformations. The range of intra- and inter-
dimer bending angles is also very similar to the recently
reported model protofilaments, 88–208 in this study com-
pared to 58–148 in the work of Grafmuller and Voth. Results
also suggest that GTP-tubulin dimers engage in relatively
stronger interdimer interactions along the filament. The
larger strength of GTP-tubulin longitudinal contacts agrees
well with their larger bending stiffness in protofilaments.
This finding is also consistent with in vitro studies, which
implicate GTP-tubulin to be better assembly candidate than
GDP-tubulin. Lastly, we show that the tubulin C-terminal
domain and the nucleotide state are closely linked with the
GTP state preventing the outward curving of C-domain.
However, this remains a testable hypothesis for future
research. The study also provides a platform for evaluating
the empirical force fields to capture the large-scale changes in
complex biological systems, such as the one investigated
here. The similar changes observed in this study using
AMBER parameters and the previous studies using
CHARMM parameters are encouraging.
The accuracy of the model tubulin structures was also
examined by performing a second control simulation. In the
model GTP-tubulin structure, the GTP ligand was switched
to GDP and the evolution of the bending angle was noted. As
Supporting Information Figure S4 shows, the original GDP-
tubulin structure was reproduced and the bending angles
switched back to the GDP bent state during the course of 100
ns simulation. Similarly, the protein–protein docking results
were checked by repeating the calculations with different set
of GTP- and GDP-tubulin structures, corresponding to the
conformations of free dimers at 50, 100, 150, and 200 ns of
FIGURE 8 (a) Time evolution of the salt-bridge between
b:Glu411 in C-domain and b:Lys105 in N-domain. Locations of the
residues are highlighted in the inset. (b) The distribution of angles
constituted by the interfacial helices H3’ and H11. Color scheme—
Red: GTP-tubulin; green: GDP-tubulin.
Nucleotide-Dependent Changes in Free Dimers to Tubulin Assembly 289
Biopolymers
simulation time. The dimer–dimer complexes were found to
be consistently bent in both states, even though the GTP-
tubulin complexes were relatively stronger.
MATERIALS AND METHODSThe coordinates for missing residues a:1, b:1, and a:35-60 in the
crystal structure were modeled using the InsightII graphics pack-
age.53 The hydrogens for heavy atoms were added by leap program
in Amber 9.0 package. Added hydrogens were energy minimized for
2000 steps using the conjugate gradient algorithm. The protonation
states of histidines—HID or HIE—were determined by the local
hydrogen bonding network using WHATIF program.54 A set of par-
tial atomic charges for GDP and GTP was obtained via quantum
electronic structure calculations. Using the Gaussian 03 program
with the 6-311G* basis set, we performed a Hartree-Fock geometry
optimization procedure. The atom-centered RESP charges were
determined via fits to the electrostatic potentials obtained from the
calculated wave functions. The missing interaction parameters in
the nucleotides were generated using antechamber tool in Amber.
After relaxing the added atoms in gas phase, the structure was
solvated in an octahedral periodic box of explicit water with water
molecules extending 12 A outside the protein on all sides. The trans-
ferable intermolecular potential three point (TIP3P) model was
chosen to describe the water molecules. To neutralize the system
and to maintain an ionic strength of 140 mM, 125 potassium and
89 chloride ions were incorporated. The system thus contained a
total of 34994 particles. Particle-Mesh Ewald summation55 with a 10
A cutoff was used to treat the long-range electrostatics. Noting that
the crystal structure used to initiate the MD simulations was deter-
mined at low resolution (e.g., in 1JFF, 15% of the rotamer and 10%
of the backbone torsion angles were flagged as outliers), an exten-
sive set of minimization and thermalization of the engineered struc-
ture was performed to allow the system to remediate the bad geom-
etry and to relax from its lattice-constrained conformation. For this,
a further 2000 steps of conjugate gradient minimization was per-
formed followed by successive heating to 310K with an temperature
increment of 25K and maintaining Ca restraints for a total duration
of 5 ns. The resulting structure was further minimized after remov-
ing the restraints and heated to 310K in 10 steps of 1 ns each. Then,
the system was equilibrated for 20 ns in NPT ensemble at 1 atm.
During this period, the potential energy of the system was seen to
be converging (Supporting Information Figure S5). The resulting
structure, thus, produces us a reliable starting model for the free
GDP-tubulin dimer. This structure was (i) further simulated to gen-
erate the 250 ns production data for GDP-tubulin system and (ii)
further engineered to prepare the starting conformation for a-GTP-
b-GTP-tubulin simulation, as the following.
In the equilibrated structure of GDP-tubulin, a c phosphate was
introduced to the bound GDP at E site to convert it to GTP. Subse-
quently, one Mg21 ion was added and the number of KCl was
adjusted to maintain the ionic strength. This new system was then
equilibrated for 20 ns following the exact procedure as described
above, which produces the starting model for free GTP-tubulin. The
simulation was extended until a trajectory of 250 ns length in NPT
ensemble was achieved. As a control, the taxol-bound tubulin crys-
tal structure was also simulated for 250 ns. The coordinate files of
the time-averaged GTP- and GDP-tubulin dimers are provided as
Supporting Information. All simulations were performed using the
NAMD package with Amber ff99SB force field56 on 64 processors of
an Infiniband Xeon E5472 linux cluster.
Models of the tubulin dimer–dimer complexes were obtained by
performing protein–protein docking calculations, using HAD-
DOCK.45 HADDOCK is a high ambiguity driven protein–protein
docking program that makes use of experimental data to search the
conformational space efficiently. The experimental information on
the interacting residues is introduced as AIRs, defined as an ambig-
uous intermolecular distance (diAB) with a maximum value of 3 A
between any atom m of an active residue i of protein A (miA) and
any atom n of both active and passive residues k of protein B (nkB)
and vice versa. The residues which are directly involved in binding
are termed active residues and the nearest neighbors are passive resi-
dues. The effective distance deffiAB for each restraint is calculated as:
deffiAB ¼
� XNatoms
miA¼1
XN Bres
k¼1
XNatoms
nkB¼1
1
d6miAnkB
��16
where Natoms indicates all atoms of a given residue and NBres the sum
of active and passive residues for the second protein. Thus, AIRs ena-
ble the search through all possible configurations around the interact-
ing site to predict the most favorable interacting pair of amino acids.
The docking protocol consists of three stages: (i) randomization of
orientations and rigid body energy minimization, (ii) semirigid simu-
lated annealing (SA) in torsion angle space, and (iii) final refinement
in Cartesian space with explicit solvent. After calculation, the struc-
tures are ranked according to their intermolecular energy which is
sum of electrostatic, van der Waals, and AIR energy terms.
In this study, the active residues for dimer–dimer docking were
chosen based on the electron crystallographic data.57 Passive resi-
dues were chosen as the nearest neighbors of the active residues
with [50% solvent accessibility, which also include the nucleotides
and the Mg21 ion. In each case, 1000 initial complex structures
were generated by randomizing the orientations of two partner
proteins whose translational and rotational movements were kept
completely free. The structures were subjected to rigid body energy
minimization, and the best 200 solutions were selected for SA
refinements. The resultant structures were classified into four clus-
ters. The lowest energy complex with correct longitudinal pitch
from GTP and GDP-tubulin docking was subjected to 50 ns MD
simulation for final refinement.
The computer resources of Computer Centre, IIT Madras are gratefully
acknowledged. KN acknowledges CSIR for the research fellowship.
REFERENCES1. Lowe, J.; Li, H.; Downing, K. H.; Nogales, E. J Mol Biol 2001,
313, 1045–1057.
2. Nogales, E.; Wolf, S. G.; Downing, K. H. Nature 1998, 391, 199–203.
3. Howard, J.; Hyman, A. A. Nature 2003, 422, 753–758.
4. Akhmanova, A.; Steinmetz, M. O. Nat Rev Mol Cell Biol 2008,
9, 309–322.
5. Valiron, O.; Caudron, N.; Job, D. Cell Mol Life Sci 2001, 58,
2069–2084.
6. Job, D.; Valiron, O.; Oakley, B. Curr Opin Biol 2003, 15,
111–117.
290 Natarajan, Mohan, and Senapati
Biopolymers
7. Ravelli, R. B. G.; Gigant, B.; Curmi, P. A.; Jourdain, I.; Sobel, S.;
Knossow, M. Nature 2004, 428, 198–202.
8. Nettles, J. H.; Li, H.; Cornett, B.; Krahn, J. M.; Snyder, J. P.;
Downing, K. H. Science 2004, 305, 866–869.
9. Gigant, B.; Wang, C. G.; Ravelli, R. B. G.; Roussi, F.; Steinmetz,
M. O.; Curmi, P. A.; Sobel, A.; Knossow, M. Nature 2005, 435,
519–522.
10. Cormier, A.; Marchand, M.; Ravelli, R. B. G.; Knossow, M.;
Gigant, B. Embo Rep 2008, 9, 1101–1106.
11. Dorleans, A.; Gigant, B.; Ravelli, R. B. G.; Mailliet, P.; Mikol, V.;
Knossow, M. Proc Natl Acad Sci USA 2009, 106, 13775–13779.
12. Wang, C.; Cormier, A.; Gigant, B.; Knossow, M. Biochemistry
2007, 46, 10595–10602.
13. Barbier, P.; Dorleans, A.; Devred, F.; Sanz, L.; Allegro, D.;
Alfonso, C.; Knossow, M.; Peyrot, V.; Andreu, J. M. J Biol Chem
2010, 285, 31672–31681.
14. Keskin, O.; Durell, S. R.; Bahar, I.; Jernigan, R. L.; Covell, D. G.
Biophys J 2002, 83, 663–680.
15. Sept, D.; Baker, N. A.; McCammon, J. A. Protein Sci 2003, 12,
2257–2261.
16. VanBuren, V.; Cassimeris, L.; Odde, D. J. Biophys J 2005, 89,
2911–2916.
17. Drabik, P.; Gusarov, S.; Kovalenko, A. Biophys J 2007, 92, 394–403.
18. Dima, R. I.; Joshi, H. Proc Natl Acad Sci USA 2008, 105, 15743–
15748.
19. Wu, Z.; Wang, H.; Mu, W.; Ouyang, Z.; Nogales, E.; Xing, J.
PLoS ONE 2009, 4, e7291.
20. Deriu, M. A.; Soncini, M.; Orsi, M.; Patel, M.; Essex, J. W.;
Montevecchi, F. M.; Redaelli, A. Biophys J 2010, 99, 2190–2199.
21. Gebremichael, Y.; Chu, J. W.; Voth, G. A. Biophys J 2008, 95,
2487–2499.
22. Bennett, M. J.; Chik, J. K.; Slyz, G. W.; Luchko, T.; Tusynnski, J.;
Sackett, D. L.; Schriemer, D. C. Biochemistry 2009, 48, 4858–4870.
23. Grafmuller, A.; Voth, G. A. Structure 2011, 19, 409–417.
24. Mitra, A.; Sept, D. Biophys J 2008, 95, 3252–3258.
25. Mitra, A.; Sept, D. J Med Chem 2006, 49, 5226–5231.
26. Mitra, A.; Sept, D. Biochemistry 2004, 43, 13955–13962.
27. Pieraccini, S.; Saladino, G.; Cappelletti, G.; Cartelli, D.; Frances-
cato, P.; Speranza, G.; Manitto, P.; Sironi, M. Nat Chem 2009, 1,
642–648.
28. Jiang, H.; Jiang, L.; Posner, J. D.; Vogt, B. D. Comput Mech
2008, 42, 607–618.
29. Sept, D.; MacKintosh, F. C. Phys Rev Lett 2010, 104, 018101.
30. Wells, D. B.; Aksimenteiv, A. Biophys J 2010, 99, 629–637.
31. Nawrotek, A.; Knossow, M.; Gigant, B. J Mol Biol 2011, 412,
35–42.
32. Natarajan, K.; Senapati, S. PLoS ONE 2012, 7, e42351.
33. Amos, L. A.; Lowe, J. Chem Biol 1999, 6, R65–R69.
34. Wang, H. W.; Nogales, E. Nature 2005, 435, 911–915.
35. Muller-Reichert, T.; Chretien, D.; Severin, F.; Hyman, A. A. Proc
Natl Acad Sci USA 1998, 95, 3661–3666.
36. Nogales, E.; Wang, H. Curr Opin Cell Biol 2006, 18, 179–184.
37. Elie-Caille, C.; Severin, F.; Helenius, J.; Howard, J.; Muller, D. J.;
Hyman, A. A. Curr Biol 2007, 17, 1765–1770.
38. Oliva, M. A.; Cordell, S. C.; Lowe, J. Nat Struc Mol Biol 2004,
12, 1243–1250.
39. Schlieper, D.; Oliva, M. A.; Andreu, J. M.; Lowe, J. Proc Natl
Acad Sci USA 2005, 102, 9170–9175.
40. Aldaz, H.; Rice, L. M.; Stearns, T.; Agard, D. A. Nature 2005,
435, 523–527.
41. Buey, R. M.; Diaz, J. F.; Andreu, J. M. Biochemistry 2006, 45,
5933–5938.
42. Oliva, M. A.; Trambailo, D.; Lowe, J. J Mol Biol 2007, 373,
1229–1242.
43. Rice, L. M.; Montabana, E. A.; Agard, D. A. Proc Natl Acad Sci
USA 2008, 105, 5378–5383.
44. Mozziconacci, J.; Sandblad, L.; Wachsmuth, M.; Brunner, D.;
Karsenti, E. PLoS ONE 2008, 3, e3821.
45. Dominguez, C.; Boelens, R.; Bonvin, A. M. J. J. J Am Chem Soc
2003, 125, 1731–1737.
46. Sobolev, V.; Sorokine, A.; Prilusky, J.; Abola, E. E.; Edelman, M.
Bioinformatics 1999, 15, 327–332.
47. Lucho, T.; Huzil, J. T.; Stepanova, M.; Tuszynski, J. Biophys J
2008, 94, 1971–1982.
48. Al-Bassam, J.; Ozer, R. S.; Safer, D.; Halpain, S.; Milligan, R. A. J
Cell Biol 2002, 157, 1187–1196.
49. Lefevre, J.; Chernov, K. G.; Joshi, V.; Delga, S.; Toma, F.; Pastre,
D.; Curmi, P. A.; Savarin, P. J Biol Chem 2011, 286, 3065–3078.
50. Padila, R.; Otin, C. L.; Serrano, L.; Avila, J. FEBS Lett 1993, 325,
173–176.
51. Popodi, E. M.; Hoyle, H. D.; Turner, F. R.; Raff, E. C. Cell Mot
Cytoskel 2005, 62, 48–64.
52. Zanic, M.; Stear, J. H.; Hyman, A. A.; Howard, J. PLoS ONE
2009, 4, e7585.
53. Accelrys Inc. Insight II 1996; Accelrys Inc.: San Diego, CA,
1996.
54. Vriend, G. J Mol Graph 1990, 8, 52–56.
55. Essmann, U.; Perera, L.; Berkowitz, M. L.; Darden, T.; Lee, H.;
Pederson, L. G. J Chem Phys 1995, 103, 8577–8593.
56. Hornak, V.; Abel, R.; Okur, A.; Strockbine, B.; Roitberg, A.;
Simmerling, C. Proteins 2006, 65, 712–725.
57. Nogales, E.; Whittaker, M.; Milligan, R. A.; Downing, K. H. Cell
1999, 96, 79–88.
Reviewing Editor: J. Andrew McCammon
Nucleotide-Dependent Changes in Free Dimers to Tubulin Assembly 291
Biopolymers