Article

Structure of Shroom domain 2 reveals a three-segmented coiled-coil required for dimerization, Rock binding, and apical constriction

Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA 15260, USA.
Molecular biology of the cell (Impact Factor: 4.47). 04/2012; 23(11):2131-42. DOI: 10.1091/mbc.E11-11-0937
Source: PubMed

ABSTRACT

Shroom (Shrm) proteins are essential regulators of cell shape and tissue morpho-logy during animal development that function by interacting directly with the coiled-coil region of Rho kinase (Rock). The Shrm-Rock interaction is sufficient to direct Rock subcellular localization and the subsequent assembly of contractile actomyosin networks in defined subcellular locales. However, it is unclear how the Shrm-Rock interaction is regulated at the molecular level. To begin investigating this issue, we present the structure of Shrm domain 2 (SD2), which mediates the interaction with Rock and is required for Shrm function. SD2 is a unique three-segmented dimer with internal symmetry, and we identify conserved residues on the surface and within the dimerization interface that are required for the Rock-Shrm interaction and Shrm activity in vivo. We further show that these residues are critical in both vertebrate and invertebrate Shroom proteins, indicating that the Shrm-Rock signaling module has been functionally and molecularly conserved. The structure and biochemical analysis of Shrm SD2 indicate that it is distinct from other Rock activators such as RhoA and establishes a new paradigm for the Rock-mediated assembly of contractile actomyosin networks.

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Available from: Michael A Trakselis
Volume 23 June 1, 2012 2131
M BoC | ARTICLE
Structure of Shroom domain 2 reveals
a three-segmented coiled-coil required for
dimerization, Rock binding, and apical
constriction
Swarna Mohan
a
, Ryan Rizaldy
a
, Debamitra Das
a
, Robert J. Bauer
b
, Annie Heroux
c
,
Michael A. Trakselis
b
, Jeffrey D. Hildebrand
a
, and Andrew P. VanDemark
a
a
Department of Biological Sciences and
b
Department of Chemistry, University of Pittsburgh, Pittsburgh, PA 15260;
c
Department of Biology, Brookhaven National Laboratory, Upton, NY 11973
ABSTRACT Shroom (Shrm) proteins are essential regulators of cell shape and tissue morpho-
logy during animal development that function by interacting directly with the coiled-coil re-
gion of Rho kinase (Rock). The Shrm–Rock interaction is sufficient to direct Rock subcellular
localization and the subsequent assembly of contractile actomyosin networks in defined sub-
cellular locales. However, it is unclear how the Shrm–Rock interaction is regulated at the
molecular level. To begin investigating this issue, we present the structure of Shrm domain
2 (SD2), which mediates the interaction with Rock and is required for Shrm function. SD2 is a
unique three-segmented dimer with internal symmetry, and we identify conserved residues
on the surface and within the dimerization interface that are required for the Rock–Shrm in-
teraction and Shrm activity in vivo. We further show that these residues are critical in both
vertebrate and invertebrate Shroom proteins, indicating that the Shrm–Rock signaling mod-
ule has been functionally and molecularly conserved. The structure and biochemical analysis
of Shrm SD2 indicate that it is distinct from other Rock activators such as RhoA and estab-
lishes a new paradigm for the Rock-mediated assembly of contractile actomyosin networks.
INTRODUCTION
Members of the Shroom (Shrm) family of cytoskeletal adaptor
proteins bind to Rho-associated coiled-coil kinase (Rock) and are
important determinants of cytoskeletal organization, cellular be-
havior, and tissue shape (Hildebrand and Soriano, 1999; Fairbank
et al., 2006; Hagens et al., 2006b; Nishimura and Takeichi, 2008;
Taylor et al., 2008; Lee et al., 2009; Chung et al., 2010; Plageman
et al., 2010). In vertebrates, the Shrm family consists of four mem-
bers, Shrm1–4 (Hagens et al., 2006a), and many of these have
been implicated in the morphogenesis of cells and tissues, in-
cluding the neural tube (Hildebrand and Soriano, 1999), the eye
(Lee et al., 2009; Plageman et al., 2010), vasculature (Farber
et al., 2011), neurons (Taylor et al., 2008), and intestines (Chung
et al., 2010; Plageman et al., 2011). Shrm family members have
also been implicated in X-linked mental retardation (Hagens
et al., 2006b) and renal function (Kottgen et al., 2009) in humans.
All Shrm proteins tested control cell morphology and tissue archi-
tecture by regulating the subcellular distribution of actomyosin
networks and use these to elicit apical constriction or cortical
contractility (Hildebrand, 2005). Shrm proteins are also found in
most invertebrates, and analysis of Drosophila Shrm (dShrm) sug-
gests that the principal functions of these proteins are conserved
Monitoring Editor
Benjamin Margolis
University of Michigan
Medical School
Received: Nov 22, 2011
Revised: Feb 23, 2012
Accepted: Mar 29, 2012
This article was published online ahead of print in MBoC in Press (http://www
.molbiolcell.org/cgi/doi/10.1091/mbc.E11-11-0937) on April 4, 2012.
S.M., J.D.H., and A.P.V. conceived and designed the study. S.M. carried out all
aspects of crystallization and structure determination, except data collection at
the National Synchrotron Light Source at Brookhaven National Laboratory, which
was performed by A.H. R.R. and S.M. performed the biochemical analyses. D.D.
performed the cell-based apical constriction assays. R.J.B. and M.A.T. performed
and analyzed the fluorescence energy transfer assays. A.P.V. and J.D.H. wrote the
manuscript with input from S.M. and M.A.T.
The authors declare that they have no conflict of interest.
Address correspondence to: Andrew P. VanDemark (andyv@pitt.edu), Jeffrey D.
Hildebrand (jeffh@pitt.edu).
© 2012 Mohan et al. This article is distributed by The American Society for Cell
Biology under license from the author(s). Two months after publication it is avail-
able to the public under an Attribution–Noncommercial–Share Alike 3.0 Unported
Creative Commons License (http://creativecommons.org/licenses/by-nc-sa/3.0).
“ASCB
®
,” “The American Society for Cell Biology
®
,” and “Molecular Biology of
the Cell
®
” are registered trademarks of The American Society of Cell Biology.
Abbreviations used: HD, homodimerization; SBD, Shroom-binding domain; SC,
surface cluster; SD2, Shroom domain 2; Shrm, Shroom.
Page 1
2132 | S. Mohan et al. Molecular Biology of the Cell
only minor disorder observed at the termini of each chain. The SD2
dimer adopts a highly unusual fold consisting of three antiparallel
coiled-coil segments (Figure 1B). Each monomer contains three he-
lices, with the B helix being roughly twice the length of the A and C
helices. The B helices wrap around each other to form a “body”
segment of 85 residues, whereas the A and C helices pair to form
45-residue “arm” segments (Figure 1B and Supplemental Figure
S1). Within both the arm and body segments, coiled-coil interac-
tions establish an extensive dimer interface, burying 4577 Å
2
of sur-
face area. This interface contains many conserved leucine and iso-
leucine residues, making interactions within the dimer interface
reminiscent of leucine-zipper domains. In contrast to Shrm SD2, leu-
cine zippers are most often parallel dimers; however, we note that
the structural database contains a large and diverse collection of
coiled-coil–containing proteins in both parallel and antiparallel ar-
rangements. To confirm that SD2 forms a dimer in solution, we
(Dietz et al., 2006; Bolinger et al., 2010). The activity of all Shrm
proteins is contingent upon proper subcellular localization and
their ability to bind Rock (Haigo et al., 2003; Hildebrand, 2005;
Plageman et al., 2010). The Shrm–Rock interaction is mediated
by the highly conserved Shrm domain 2 (SD2), located at the
C-termini of all Shrm proteins (Hildebrand, 2005; Dietz et al.,
2006).
Myosin II and the actin cytoskeleton are universally used by cells
to control shape and behavior in response to environmental stimuli
during a wide range of biological processes. The activity of myosin
II is tightly controlled through phosphorylation of the associated
myosin regulatory light chains by a number of serine/threonine ki-
nases and phosphatases (Ikebe et al., 1988; Moussavi et al., 1993).
One of these kinases is Rock (Amano et al., 1996a; Ishizaki et al.,
1996), which has been shown to regulate myosin II activity directly
by phosphorylating Ser-19 of the myosin light chain and indirectly
by inhibiting the activity of the myosin phosphatase (Amano et al.,
1996a; Kimura et al., 1996; Kawano et al., 1999). The activity of Rock
appears to be tightly controlled via several mechanisms. Primary
among these is relief of intramolecular inhibition of the kinase do-
main by its C-terminus. This is typically achieved by the binding of
GTP-bound RhoA to the Rho-binding domain located within the
coiled-coil region of Rock (Ishizaki et al., 1996; Matsui et al., 1996).
It is predicted that RhoA binding causes a conformation change
within Rock that displaces the C-terminus from the kinase domain
and allows for catalytic activity (Amano et al., 1996b, 1999). SD2 of
Shrm has also been shown to interact with the coiled-coil region of
Rock but at a location that is distinct from the Rho-binding domain
(Nishimura and Takeichi, 2008; Taylor et al., 2008; Bolinger et al.,
2010; Farber et al., 2011).
Although structures have been determined for many portions of
Rock, including the kinase domain (Jacobs et al., 2006; Yamaguchi
et al., 2006a, 2006b; Komander et al., 2008), PH domain (Wen et al.,
2008), portions of the coiled-coil domain (Shimizu et al., 2003; Tu
et al., 2011), and a RhoA:Rho-binding domain complex (Dvorsky
et al., 2004), there is no structural information on the Shrm-binding
domain of Rock and no structural information for any portion of
Shrm. Consequently, there is little information regarding the mo-
lecular details of the Shrm–Rock interaction or how Shrm binding
affects the activation status of Rock. Here, we take a structural ap-
proach to gain molecular and mechanistic insight into SD2 of Shrm
and its interaction with Rock.
RESULTS
SD2 adopts an extended three-segmented coiled-coil
To understand the molecular basis for Shrm-mediated regulation of
actomyosin contractility, we initiated a structural analysis of Shrm
proteins. These studies focused on the C-terminal SD2 since it is the
most highly conserved domain found in all Shrm family members
and is both necessary and sufficient for activating actomyosin con-
tractility (Hildebrand, 2005). Limited proteolysis of various SD2-con-
taining protein fragments derived from mouse Shrm3 indicates the
presence of a stable “core” of 180 residues located at the C-termi-
nus of SD2. We used these data to guide the design of SD2 expres-
sion constructs from several different Shrm proteins. We were able
to obtain and optimize crystals from dShrm containing amino acid
residues 1393–1576 (Figure 1A) and determine its structure using
the SAD method with selenomethionine (SeMET)-substituted crys-
tals (see Material and Methods and Table 1 for a complete descrip-
tion of the structure determination procedure).
The structure is refined at 2.7-Å resolution with an R
free
value of
27.4%. The asymmetric unit contains a complete SD2 dimer, with
SeMET (SAD) Native
Data collection
Space group P2
1
2
1
2 P2
1
2
1
2
Cell dimensions
a (Å) 72.2 72.8
b (Å) 84.9 85.6
c (Å) 93.0 93.0
Resolution (Å) 30.0–3.5
(3.56–3.50)
50.0–2.7
(2.75–2.70)
Unique reflections 7573 16,446
R
merge
8.5 (8.2) 6.9 (46.3)
I/σI
42.1 (34.1) 34.2 (3.5)
Completeness (%) 99.3 (100) 99.9 (99.9)
Redundancy 10.0 (10.7) 8.5 (8.1)
Refinement
Resolution (Å) 47.0–2.7
R
work
/R
free
22.78/28.38
Number of protein atoms 2749
Root mean square
deviations
Bond lengths (Å) 0.10
Bond angles (deg) 0.600
Average isotropic B
values (Å
2
)
75.9
Ramachandran statistics
Outliers 0
Allowed 0
Favored 100
Values in parentheses correspond to those in the outer resolution shell.
R
merge
= (|(Σ I <I>)|)/(Σ I), where <I> is the average intensity of multiple mea-
surements.
R
work
= Σ
hkl
F
obs
(hkl) F
calc
(hkl)/Σ
hkl
|F
obs
(hkl)|.
R
free
= cross-validation R factor for 7.3% of the reflections against which the
model was not refined.
TABLE 1: Data collection and refinement statistics for dShrm SD2.
Page 2
Volume 23 June 1, 2012 Structure of Shroom domain 2 | 2133
acterized SD2’s solution state using gel filtration (Figure 1D). We
observe two species in this assay: a larger dimeric species that was
used for crystallization and a minor peak containing 9% of the peak
area. These data indicate that the dimeric species we observe in the
crystal is the predominant species in solution.
There are notable regions of both symmetry and asymmetry
within SD2. The molecule is internally symmetric, with the left and
right half-dimers exhibiting near structural identity (root mean square
deviation of 0.6 Å over 174 Cα atoms; Figure 1B and Supplemental
treated purified SD2 with the chemical cross-linker glutaraldehyde
and resolved the resulting species on SDS–PAGE (Figure 1C). These
assays indicate that we can readily detect a dimeric SD2 species in
solution (Figure 1C). In fact in the absence of cross-linker, a small
dimeric fraction is still observed in the SDS–PAGE gel, indicating the
strength of interaction in the coiled-coil. In this assay, we can also
detect tetrameric and other higher-order species that appear to be
formed by spurious cross-linking between SD2 dimers. Because this
technique is not quantitative (Trakselis et al., 2005), we further char-
FIGURE 1: Structure of the dShrm SD2 dimer. (A) Domain organization for the Shroom proteins used in this study.
The predicted secondary structure for the canonical SD2 and the actual secondary structure and the location of
relevant features from the crystallized fragment are shown. (B) Ribbon diagram of the dShrm SD2 dimer. The body
segment, two arm segments, and the symmetry point locations are indicated. (C) Chemical cross-linking of dShrm
SD2. Purified dShrm SD2 was incubated with 0.009% glutaraldehyde over the indicated time period and the resulting
species separated by SDS–PAGE. (D) Gel filtration profile of wild-type dShrm SD2. Two species are observed, and the
relative peak area from each is indicated. Fractions collected during this run were analyzed by SDS–PAGE and
indicated below the trace.
Page 3
2134 | S. Mohan et al. Molecular Biology of the Cell
which is similar in length to the SD2s that are shown to cause apical
constriction (Hildebrand, 2005; Dietz et al., 2006; Figure 2A). For
Rock, we used amino acids 707–946 of human Rock1 and amino
acids 724–938 of Drosophila Rock. These sequences were chosen
based on the previously described Shrm-binding sequences
(Nishimura and Takeichi, 2008; Bolinger et al., 2010; Farber et al.,
2011), sequence conservation, and predicted secondary structure.
We refer to these regions of hRock and dRock as the Shrm-binding
domain (SBD). Because this sequence is 95% identical between
mouse and human Rock, we predicted that human Rock should
bind equally well to mouse Shrm3. In this assay, all three SD2 frag-
ments are able to bind Rock, indicating that the crystallized frag-
ment of dShrm contains a Rock-binding surface and that this surface
is likely conserved in all SD2s. To follow up on these findings, we
tested by native gel electrophoresis whether Rock and Shrm could
form a stable complex (Figure 2B). Results indicate that the Shrm–
Rock interaction is stable, saturable, and stoichiometric. Finally, to
demonstrate that the SD2 regions of mShrm3 and dShrm exhibit
equivalent functions in vivo, we tested their ability to mediate apical
constriction in cultured Madin–Darby canine kidney (MDCK) cells.
The C-terminal regions of dShrm (residues 1144–1576) and mShrm3
(residues 1372–1976), containing the SD2 motifs, were expressed as
chimeric fusion proteins consisting of the apically targeted trans-
membrane protein endolyn (Hildebrand, 2005). We also expressed
a fusion protein containing mShrm3 1372–1562 (lacking the SD2) as
a negative control. MDCK cells transiently transfected with these
expression vectors were grown on Transwell filters and stained to
detect the tight-junction marker ZO1 and the ectopically expressed
endolyn–Shrm protein. The distribution of ZO-1 (red) indicates the
Figure S2). We term the point separating the left and right halves of
the dimer the symmetry point. Of interest, there is a twist within the
dimer such that the right and left arms are rotated 60° relative to
the long axis of the body segment, which introduces an element of
asymmetry into the overall structure (Figure 1B and Supplemental
Figure S2). Structural homology searches failed to identify any struc-
tures whose similarity with Shrm extends beyond a single coiled-coil
segment, indicating that the structure we observe may be unique.
More important, the structure of SD2 is distinct from that of RhoA,
the other known activator of Rock that binds to the coiled-coil
region.
The dShrm SD2 core is sufficient for dRock binding
and apical constriction
Previous studies showed that the direct interaction between SD2 of
mouse Shrm3 (1563–1986) and the coiled-coil domain of human
Rock (698–957) is required for apical constriction (Nishimura and
Takeichi, 2008). In addition, we also showed that this interaction is
conserved in dShrm and dRock (Bolinger et al., 2010). Our structure
is missing the N-terminal 70 residues of the previously defined SD2
(Dietz et al., 2006), as these were removed to facilitate crystalliza-
tion. To demonstrate that the structure we observed still contained
the biologically relevant portion of the SD2, we examined the ability
of SD2 regions from mShrm3 and dShrm to both interact with Rock
and mediate apical constriction in a cell-based assay. To examine
the Shrm–Rock interaction, we first performed pull-down assays us-
ing histidine (His)-tagged Shrm-SD2 constructs containing the core
fragment from dShrm, the equivalent core fragment from mouse
Shrm3 (1762–1952), or a longer form of mouse Shrm3 (1543–1985),
FIGURE 2: The SD2 core is sufficient for Rock binding and apical constriction. (A) Purified His-tagged mShrm3 full SD2
(1643–1986), His-tagged mShrm3 SD2 core (1762–1952), or His-tagged dShrm SD2 core (1393–1576) was mixed with
either hRock (707–938) or dRock (724–948) as indicated and complexes detected by pull-down with Ni beads. P, pellet
fraction; S, supernatant fraction. (B) Native-PAGE of dShrm SD2 alone and mixed with increasing concentrations of
dShrm SD2. Complex formation is monitored by the formation of a slower-migrating species. (C) Endolyn-tagged Shrm
constructs were expressed in MDCK cells, and cells were stained to detect the exogenous endolyn–Shrm protein
(green) and ZO-1, a marker for tight junctions (red). Both mShrm3 SD2 and dShrm SD2 can cause apical constriction
upon being targeted to the apical membrane. Arrowheads denote cells expressing endolyn–Shrm proteins.
Page 4
Volume 23 June 1, 2012 Structure of Shroom domain 2 | 2135
was formed by both SD2 chains. Given the
large and extended dimerization interface,
we were concerned that small perturba-
tions, such as single–amino acid changes,
might not destabilize enough of the Shrm–
Shrm interface to result in measurable
changes in either dimerization or Rock bind-
ing. To avoid this potential problem, we
used sequence conservation combined with
our structural data to identify regions where
alterations within the Shrm–Shrm interface
may have the greatest impact. We identi-
fied two regions and generated multiple
substitutions to target these regions (Figure
3A and Supplemental Figure S1). We
termed these variants homodimerization
(HD) mutants. One interface mutant, HD1
(
1468
LLSL
1471
to AASA; Figure 3A), primarily
targets the body segment, whereas the sec-
ond HD mutant, HD2 (
1546
LIADARDL
1553
to
AAADARDA; Figure 3A), primarily targets
the coiled-coil within the arm segment.
These amino acid changes are also pre-
dicted to weaken contacts between the arm
and body segments but to a lesser degree.
The selected amino acids were changed to
alanine, as its high helical propensity should
minimize effects due to alterations in sec-
ondary structure. We expressed and puri-
fied these proteins and compared their elu-
tion profile in gel filtration to wild-type
protein (Figure 3B). We observe distinct
changes with both mutants; protein contain-
ing the HD1 substitution elutes in a single
broad peak distinct from both species ob-
served with the wild-type protein. HD2 has
an equally pronounced but different effect,
in which much of the dimeric peak has been
shifted into a larger, uncharacterized spe-
cies. We isolated protein corresponding to
dimer in the case of HD2 or to the majority
peak from HD1 purification (Figure 3B) and
further characterized the effect of substitu-
tions within the dimerization interface. We
first tested their ability to form homodimers
in solution by chemical cross-linking (Figure
3C). In this assay, both HD mutants exhib-
ited reduced cross-linking when compared
with wild type, indicating a change in the dimeric interface. It should
be noted that the substitutions in HD1 are more severe and perturb
dimerization to a greater extent than those substitutions in HD2. To
further confirm that our HD variants perturb the structure of SD2, we
probed their stability via limited proteolysis using the nonspecific
enzyme subtilisin A (Supplemental Figure S3). Although still readily
expressed and purified, both HD variants are more accessible to
protease, indicating a disruption of the dimerization interface. Con-
sistent with the data obtained in the cross-linking experiment de-
scribed here, HD1 appears to be more sensitive to proteolysis. We
then tested the ability of the HD mutant proteins to bind dRock by
native gel electrophoresis (Figure 3D). Neither variant is able to bind
the dRock-SBD (724–938), indicating that these substitutions alter
the positions of residues within Shrm that are required for Rock
apical boundaries, whereas Shrm (green) localization indicates that
all three fusions were expressed and targeted to the apical plasma
membrane. Cells expressing an endolyn fusion containing an intact
SD2 are able to constrict, whereas the control endolyn–Shrm3
fusion is unable to perform apical constriction (Figure 2C). Therefore
we conclude that the crystallized SD2 contains the Rock-binding
site and, when properly localized, is sufficient to mediate apical
constriction.
Perturbation of the SD2 dimerization interface inhibits
Rock binding
We next examined whether the SD2 dimerization interface was
important for Rock binding, reasoning that the extended shape
observed for SD2 made it more likely that the Rock-binding site
FIGURE 3: Mutations in the dimerization interface diminish Rock binding. (A) Ribbon diagram of
SD2 highlighting the interface mutations, HD1 (green), and HD2 (blue). Residues making
contacts with HD1 or HD2 are shown as white (chain A) or gold (chain B) sticks. (B) Gel filtration
chromatograms for wild-type and HD1 and HD2 mutant proteins. SDS–PAGE of resulting
fractions aligned to the chromatogram is shown below. (C) Chemical cross-linking of wild-type
and HD1 and HD2 mutant proteins. The indicated dShrm SD2 protein was incubated with
0.002% glutaraldehyde. Samples were taken at the indicated time points and resolved by
SDS–PAGE. (D) Native gel electrophoresis of dRock 724–938 mixed with increasing
concentrations of wild-type and HD mutant dShrm–SD2 proteins.
Page 5
2136 | S. Mohan et al. Molecular Biology of the Cell
elongated nature of the conserved clusters,
we were concerned that the in vitro binding
studies may not prove sensitive enough
to observe changes resulting from single–
amino acid changes. Therefore we de-
signed three multiresidue variants with al-
terations on the SD2 surface while avoiding
residues that could play a role in dimeriza-
tion. The surface cluster (SC) variants are
1402
KMDEL
1406
to AMDRA,
1470
SLSERLA
1476
to ALEEDLE, and
1509
LKSDIERR
1516
to AAS-
DIEDA, which for clarity are named SC1,
SC2, and SC3, respectively. The locations of
these substitutions within the SD2 surface
are indicated in Figure 4 (green residues).
The elution profiles for the surface cluster
variants were largely unchanged relative to
wild type, suggesting that these mutations
do not significantly alter the overall struc-
ture of the SD2 dimer (Supplemental Figure
S5). We tested the surface cluster variants
for their ability to bind dRock-SBD by pull-
down (Figure 4C). In this assay, His-tagged
dRock effectively precipitates wild-type
SD2 and SC1. In contrast, this interaction is
abrogated by substitutions made in SC2
and 3. We also monitored formation of a
dShrm–dRock complex by native gel elec-
trophoresis (Figure 4B). Similar to the results
with the pull-down, SC1 binds dRock-like
wild type, whereas complex formation with
SC3 is undetectable. Although we could
detect some complex formation with the
SC2 variant, binding is clearly reduced, in-
dicating that the targeted amino acids are
located within the Rock-binding surface. These data indicate that
Rock binding is most likely mediated by amino acids within the
body segment, whereas the cluster of conserved residues within
the arm is not involved. This supports the hypothesis that the Rock-
binding site is composed of residues on the surface of the SD2 di-
mer. Further, since the SC2 derivative exhibits an intermediate level
of binding, we conclude that these amino acids may lie at the pe-
riphery of the Rock binding site, whereas SC3 contains residues
that are more critical for Rock binding.
The Rock-binding interface is conserved in
vertebrate Shroom
We next tested whether the residues we show play an important
role in Shrm–Rock binding in Drosophila are conserved in verte-
brates. We noted that there was considerable sequence conserva-
tion within SD2s from various vertebrate Shrm proteins, so we chose
to examine the effect of mutations within the context of mouse
Shrm3 due to its ability to induce apical constriction in MDCK cells.
The following amino acid changes were made in mShrm3 SD2
and the subsequent proteins tested for the ability to homodimerize
and bind to the SBD of human Rock1:
1766
KKAEL
1770
to AKARA
(SC1),
1834
SLSGRLA
1840
to ALEADLE (SC2),
1878
LKENLDRR
1885
to
AAENLDDA (SC3),
1832
LLSL
1835
to AASA (HD1), and
1915
LLIEQRKL
1922
to ALIEQAKA (HD2). All of the homo dimerization and surface clus-
ter mutations were generated in a plasmid encoding glutathione
S-transferase (GST)tagged mShrm3 SD2. Purified proteins were
first tested for the ability to bind the hRock SBD (Figure 5A). In this
binding. Taken together, these data indicate that mutations that
perturb the Shrm–Shrm interface have a dramatic effect on Rock
binding and suggest that the Rock-binding site on Shrm is com-
posed of elements from both chains of the dimer.
A conserved Rock-binding interface on the SD2 surface
On the basis of the forgoing results, we hypothesized that we would
be able to identify patches of surface residues that are required for
binding to Rock but are not involved in dimerization. To test this, we
searched for conserved patches of amino acids on the surface on
the SD2 dimer by aligning 12 Shrm sequences from both vertebrate
and invertebrate organisms (Supplemental Figure S1). We then used
the RISLER matrix (Risler et al., 1988), as implemented in ESPRIPT
(Gouet et al., 1999), to score and map conservation onto the SD2
surface (Figure 4A). Although this domain is highly conserved
throughout its entire sequence, we identified three clusters of highly
conserved residues as candidates for the Rock-binding surface. Two
of these surfaces lie on opposite faces of the main body segment
within helix B, whereas a third surface is formed by residues within
helix A found near the end of the arm segment (Figure 4A). It should
be noted that these patches are derived from amino acids residues
on both the A and B chains, supporting the hypothesis that di-
merization may be required to form a functional binding surface.
To address the importance of these surface clusters in Rock
binding, we used the structural data to design amino acid substitu-
tions within these potentially important surfaces. Given the prepon-
derance of invariant residues, their broad distribution, and the
FIGURE 4: Conserved surfaces on SD2 are important for dRock binding. (A) Surface of SD2 with
sequence conservation mapped in shades of blue. Invariant residues within SC mutants are
shown in green. Three extended surfaces with high sequence conservation are outlined in yellow
for clarity. (B) Native gel electrophoresis of dRock mixed with the indicated SD2 mutants.
(C) Pull-down assay using His-dRock and indicated SD2 mutants.
Page 6
Volume 23 June 1, 2012 Structure of Shroom domain 2 | 2137
ter mutations had no effect on binding to SD2. It should be noted
that the surface cluster variant 1 bound with slightly reduced effi-
ciency. On the basis of these data, we conclude that the Rock-bind-
ing interface identified in Drosophila is largely conserved in the
mouse proteins and that this Shrm–Rock binding module has been
conserved across animal evolution at both the molecular and func-
tional levels.
The Rock-binding surface is required for apical constriction
Our previous work showed that the SD2 motif of Shrm3 is both
necessary and sufficient to cause apical constriction of polarized
assay, we could not detect binding of either of the homodimeriza-
tion variants to the Rock SBD. For the surface cluster derivatives,
binding of variant 1 to Rock was unaltered, whereas surface cluster
variants 2 and 3 were incapable of binding Rock. These results are
consistent with those obtained using the Drosophila proteins but
suggest that the surface cluster 2 region of mouse Shrm3 may play
a more significant role in binding to Rock. We next assayed the abil-
ity of the surface cluster and homodimerization variants to form
homodimers with an untagged, wild-type mShrm3-SD2 (Figure 5B).
As expected from our studies with dShrm, the homodimerization
mutations severely impaired dimerization, whereas the surface clus-
FIGURE 5: The Rock-binding interface is conserved in vertebrate Shroom. (A) Wild-type and mutant GST-tagged mouse
Shrm3 SD2 proteins were mixed with untagged hRock as indicated and complexes detected by pull-down with
glutathione resin followed by SDS–PAGE and Coomassie staining. (B) The ability of GST-tagged interface or surface
cluster mutants to bind untagged mShrm3 SD2 was tested by a pull-down assay. (C) Wild-type and SD2 mutant versions
of endolyn-tagged mShrm3 were expressed in MDCK cells and cells stained to detect Shrm3, ZO-1, and ppMLC. Only
the wild type and the SD1 variant induce apical constriction and recruitment of active myosin II when targeted to the
apical membrane. Transfected cells are denoted by arrowheads; scale bar, 20 μm.
Page 7
2138 | S. Mohan et al. Molecular Biology of the Cell
no enrichment of active myosin II. These data suggest that in vivo,
the SD2 motif must retain the ability to both dimerize and bind Rock
in order to trigger apical constriction and that Shrm3-mediated api-
cal contraction is dependent on the activity of both Rock and
myosin.
Characterizing the Shrm–Rock complex
In an effort to elucidate the molecular details of the Shrm–Rock com-
plex, we first used fluorescence energy transfer (FRET) experiments
to detect and quantify the interaction between dShrm and dRock
SBD. Because the precise binding interface between dRock and
dShrm is unknown, we labeled dRock with Cy5 at its N-terminus,
whereas dShrm SD2 was labeled with Cy3 at a single cysteine (C1428)
not believed to be located within the Rock-binding interface. There
are two endogenous cysteines within this fragment of dShrm, so a
conservative mutant of dShrm (C1533S) was generated for this assay
to ensure labeling at a single position. Titration of dShrm with dRock
resulted in a decrease in donor emission and increase in acceptor
emission consistent with an increase in FRET due to a binding inter-
action (Figure 6A). Assuming a single-binding mode for this interac-
tion, we calculate the equilibrium K
d
to be 0.58 ± 0.07 μM (Figure 6B).
This affinity is comparable to that of RhoA, which has a reported K
d
of 0.13 μM (Blumenstein and Ahmadian, 2004).
epithelial cells when targeted to the apical domain of the cell
(Hildebrand, 2005). To test whether alterations to the dimerization
interface or the Rock-binding surface affect the ability of the Shrm3
SD2 to induce apical constriction, we introduced our homodimeriza-
tion and surface cluster amino acid substitutions into the endolyn–
mShrm3 chimeric protein. All of the endolyn–Shrm3 variants are
expressed at equal levels and are efficiently targeted to the apical
surface (Figure 5C, arrowheads). Consistent with the in vitro binding
results, we observed that only the wild type and the surface cluster
1 variant retained the capacity to trigger apical constriction in cells.
To determine whether the various homodimerization and surface
cluster mutants were capable of activating the Rock–myosin II path-
way, we stained cells expressing each of the SD2 mutants to detect
the myosin light chain (MLC) phosphorylated at Thr-18 and Ser-19
(ppMLC), a readout of active myosin II. Consistent with the in vitro
binding assay and the foregoing results, only wild type and the
surface cluster variant 1 of endolyn–Shrm3 showed recruitment of
activated myosin II to the constricted apical surface (Figure 5C). By
measuring the increase in apical fluorescence relative to the de-
crease in apical area, we estimate that there was an approximate
1.4- to 1.8-fold increase in the amount of apically localized active
myosin II. In contrast, neither homodimerization variant nor the sur-
face cluster variants 2 or 3 caused apical constriction, and there was
FIGURE 6: Characterizing the Shrm–Rock complex. (A) FRET titration of Cy5-labeled dRock into 50 nM Cy3-labeled
dShrm showing donor quenching and acceptor sensitization for representative concentrations. (B) Donor quenching
plotted as a function of Rock concentration and fitted to a single-binding mode to give a K
d
value of 0.58 ± 0.07 μM.
The error bars show the SE for the average of at least three independent experiments. (C) Estimation of the Shrm–Rock
complex stoichiometry. Native-PAGE stained with colloidal blue was used to identify the Shrm–Rock complex as
described earlier. Bands corresponding to the complex (denoted by the asterisk) were excised from native-PAGE,
protein eluted from the gel slice, and run on SDS–PAGE to separate the components contained within. (D) Models
describing one potential mode of interaction between Shrm SD2 and Rock formed by hinging at the symmetry point
within the observed SD2 dimer.
Page 8
Volume 23 June 1, 2012 Structure of Shroom domain 2 | 2139
dimer without a major disruption to the observed SD2 conforma-
tion. We do not favor this model, however, because it is difficult to
envision how the two Rock-binding interfaces, one in each half-
dimer, would contact the two independent Shrm-binding sites that
would be generated by the nature of the parallel coiled-coil of the
Shrm-binding domain. Instead we favor a model in which there is a
large conformational change upon Rock binding that allows the SD2
to position its half-dimers on opposite sides of the Rock coiled-coil
(Figure 6D). This would allow the two surface clusters that bind to
Rock to interact with the helices of the SBD simultaneously. A direct
observation of SD2 in other conformations or bound to Rock will be
required to address this.
Implications of the Shroom–Rock interaction
It has been shown that Shrm–Rock interactions are vital for several
developmental processes, including neural tube, lens, and gut mor-
phogenesis. There is no information about the stoichiometry or af-
finity of the complex, and it is unclear how the Shrm–Rock interac-
tion may by regulated. There are two primary models for thinking
about how Shrm may function with Rock to achieve localized activa-
tion of contractile actomyosin networks. First, Shrm binding to Rock
leads to both the redistribution of Rock and the activation of its cata-
lytic activity. Second, it is possible that Shrm binding can alter the
distribution of Rock but that additional inputs activate Rock. Our
results indicate that Shrm and Rock bind with high affinity and are
likely to form a heterotetramer in solution. On the basis of the fact
that Shrm binds to Rock in close proximity to the Rho-binding site,
it is tempting to speculate that Shrm binding activates Rock in a
manner similar to Rho. However, additional structural studies and
kinetic assays will be required to verify this hypothesis.
Genetic and cell-based approaches demonstrated that the Rock–
myosin II pathway is used to control the cell behaviors that facilitate
tissue morphogenesis in animals. As a result, targeted Rock inhibi-
tion is viewed as a viable therapeutic approach for treating many
clinical conditions, including cancer (Liu et al., 2011), obesity (Hara
et al., 2011), type I diabetes (Biswas et al., 2011), pulmonary hyper-
tension (Connolly and Aaronson, 2011), and many others (reviewed
in Dong et al., 2011). The central role of Rock also makes global in-
hibition of Rock a challenge due to possible side effects. Therefore
it would be of great benefit to be able to target specific steps of
Rock activation or specific effectors of Rock. One of the ways to ac-
complish this is to understand how specific proteins interact with
Rock and elucidate the outcomes of these interactions on Rock ac-
tivity. The identification of the Shrm–Rock interaction as a distinct
module that may function independent of RhoA may provide ways
to abrogate or enhance specific arms of Rock signaling while leaving
others unperturbed.
MATERIALS AND METHODS
Protein expression and purification
Coding sequences for dShrm SD2 (residues 1393–1576) and dRock
SBD (724–938) were amplified by PCR and cloned into the bacterial
expression vector pET151-D/Topo (Invitrogen, Carlsbad, CA). Pro-
tein expression was performed in BL21(DE3) Escherichia coli cells
using ZY autoinduction media (Studier, 2005) at room temperature
for 24 h, harvested by centrifugation, and lysed via homogeniza-
tion in 25 mM Tris, pH 8.0, 500 mM NaCl, 10% glycerol, 5 mM imi-
dazole, and 5 mM β-mercaptoethanol. The lysate was cleared by
centrifugation at 100,000 × g. dSD2 was purified by nickel affinity
chromatography (Qiagen, Valencia, CA), followed by overnight di-
gestion with tobacco etch virus (TEV) protease. A second round of
nickel affinity purification was performed to remove the liberated
We next examined the stoichiometry of the dShrm–dRock com-
plex. To determine this, we mixed purified dRock SBD and dShrm
SD2 in solution to form a complex and then resolved it on a native
gel. After electrophoresis, the complexes were excised from the gel,
eluted, resolved by SDS–PAGE, and detected by Coomassie blue
staining (Figure 6C). Alternatively, complex was run on a gel filtra-
tion column and peak fractions were resolved by SDS–PAGE. The
ratio of SD2 to SBD in the complex was measured by densitometry
and corrected for the relative molecular masses of the two proteins
(Supplemental Figure S4). In all cases, isolated complexes were
composed of SD2 and SBD in an 1:1 molar ratio. Although the
possibility for a variety of higher-order species cannot be ruled out
from these data, we feel that heterodimeric and heterotetrameric
species are the most probable. This is consistent with RhoA, which
also interacts with Rock in a 1:1 molar ratio, and places important
mechanistic constraints on the complex.
DISCUSSION
Shroom domain 2 adopts a unique fold
Our studies of SD2 reveal that this motif is composed of an unusual
arrangement of three canonical coiled-coil segments. On the basis
of the structure and in vitro binding assays, we propose that two
binding surfaces within SD2 are important for Rock interaction. The
first mediates SD2 dimerization, which in turn positions conserved
residues on the SD2 surface into an orientation that is competent for
Rock binding. Conserved residues on the surface are located in
three clusters; however, only residues within the main body were
shown to play a role in Rock binding. The conserved patches within
the main body segment contain residues from both molecules of
the SD2 dimer, which may explain why dimerization is required for
Rock binding. The observed symmetry within the SD2 dimer dic-
tates that there are two independent but identical Rock-binding
sites. Of importance, any mutation that disrupts Rock binding also
abrogates Shrm-induced apical constriction in vivo.
The Shrm–Rock complex
Crystal structures of the coiled-coil portion of Rock indicate that it
exists as a dimer (Shimizu et al., 2003; Dvorsky et al., 2004; Tu et al.,
2011), and our data suggest that the Shrm–Rock complex contains
equal ratios of SD2 and SBD. Of the possible stoichiometries for the
Shrm–Rock complex, we speculate that heterodimeric or heterote-
trameric (a dimer of dimers) species are most probable, and we fa-
vor the latter for the following reasons. First, both Shrm and Rock
components are dimers in solution. Second, a Shrm–Rock heterodi-
mer would require that both the SD2 and SBD homodimers sepa-
rate before reforming the heterodimer. We predict that there would
be a large energetic barrier to this rearrangement. Third, our results
indicate that distinct surfaces are required for Rock binding and SD2
homodimerization. Finally, the crystal structure of the Rock–RhoA
complex indicates that dimerization of the Rho-binding domain is
not altered upon binding to RhoA (Dvorsky et al., 2004).
Molecular models for the Shrm–Rock complex
The dShrm SD2 structure presented here places a number of con-
straints on how it interacts with the SBD of Rock. Previous studies
showed that regions of Rock just N-terminal and C-terminal of the
Shrm-binding domain form a parallel coiled-coil dimer (Dvorsky
et al., 2004; Tu et al., 2011). On the basis of these studies, it is rea-
sonable to predict that Rock’s Shrm-binding domain also exists as a
parallel coiled-coil. If this is the case, we can envision two different
models for the Shrm–Rock interaction based on our structures. In
the first model, it is possible that the Shrm SD2 dimer binds the SBD
Page 9
2140 | S. Mohan et al. Molecular Biology of the Cell
article have been submitted to the Protein Data Bank (www.rcsb.org/
pdb/home/home.do) and assigned the identifier 3THF.
Chemical cross-linking
dShrm SD2 was incubated with the indicated concentration of glu-
taraldehyde in a reaction buffer containing 25 mM 4-(2-hydroxyethyl)-
1-piperazineethanesulfonic acid (HEPES), pH 7.5, 8% glycerol,
500 mM NaCl, and 5 mM β-mercaptoethanol, with a final dShrm
SD2 concentration of 8 μM. At each time point, 20 μl of the cross-
linking reaction was removed and the reaction stopped with 2 μl of
1.0 M Tris at pH 8.0 and the sample subjected to SDS–PAGE and
visualized using Coomassie blue staining.
Complex formation
Equal molar quantities of dShrm SD2 and dRock SBD were mixed at
a combined concentration of 2.4 mg/ml and dialyzed into 25 mM
Tris, pH 8.0, 8% glycerol, 150 mM NaCl, and 5 mM β-mercaptoethanol.
Complex was isolated using a Sephacryl S-300 gel filtration column
(GE Healthcare). The SD2–SBD complex eluted off the gel filtration
column in one peak distinct from that for SD2 or SBD alone. For
solution binding and native gel electrophoresis, a fixed concentra-
tion (5 μM) of dRock 724–938 was mixed with increasing concentra-
tion of dShrmSD2 (1–10 μM) and incubated for 1 h. Samples were
then loaded on 8% PAGE gels and resolved by electrophoresis at
4°C. Proteins were detected with Coomassie blue. For GST pull-
down assays using mShrm3, either wild-type GST-Shrm3 SD2 or SC
and HD mutant versions (spanning amino acids 1562–1986) bound
to beads were mixed with soluble, untagged mShrm3 SD2 (residues
1762–1952) or hRock1 (residues 707–946). Complexes were washed
with NETN (100 mM NaCl, 1 mM EDTA, 20 mM Tris, pH 8.0, 0.05%
NP-40), resuspended in SDS–PAGE sample buffer, resolved by SDS–
PAGE, and detected using Coomassie blue.
Apical constriction assays
MDCK cells were grown in EMEM supplemented with 10% fetal bo-
vine serum, penicillin/streptavidin, and l-glutamine. Apical constric-
tion assays using endolyn–dShrm, endolyn–Shrm3, endolyn–
mShrm3 dlSD2, or endolyn–Shrm3 harboring SC or HD were
performed and imaged as described (Hildebrand, 2005). Cells were
attained with the following antibodies: UPT132 (1:250, rabbit anti-
Shrm3; Hildebrand, 2005), rat anti-ZO1 (1:500; Chemicon, Temec-
ula, CA), and rabbit anti–pThr18/pSer19 MLC2 (1:50; Cell Signaling
Technology, Beverly, MA). Primary antibodies were detected using
Alexa 488 or 568–conjugated secondary antibodies (1:400; Invitro-
gen). Images were acquired using a Bio-Rad Radiance 2000 Laser
Scanning System (Bio-Rad, Hercules, CA) mounted on a Nikon E800
microscope (Nikon, Melville, NY) with 40× and 60× oil objectives
and processed using either ImageJ (National Institutes of Health,
Bethesda, MD) or Photoshop (Adobe, San Jose, CA). The fluores-
cence intensity of ppMLC was determined using ImageJ and was
achieved by measuring the average fluorescence intensity of a fixed
region of interest (ROI) at the apical surface of subsaturated confo-
cal images from expressing and nonexpressing cells. Fluorescence
intensity of the ROI was then corrected for the decrease in area of
apically constricted cells (n 20 cells/variant). Change in fluores-
cence intensity was then determined as the ratio of the corrected
intensity of constricted versus nonconstricted cells.
Fluorescence labeling
dShrm was labeled at the N-terminus with Alexa 594 succinimidyl
ester (Invitrogen) in amino labeling buffer (20 mM HEPES, pH 7.0,
100 mM NaCl, 8% glycerol) or at C1428 of the C1533S mutant with
His tag, TEV protease, and many nonspecific contaminants. Gel fil-
tration, using a Sephacryl S-200 gel filtration column (GE Health-
care, Piscataway, NJ), was performed, and peak fractions were con-
centrated to 9 mg/ml in 20 mM Tris, pH 8.0, 0.5 M NaCl, 8% glycerol,
and 5 mM dithiothreitol (DTT) using a Vivaspin concentrator
(Millipore, Billerica, CA) before crystallization. The purity was typi-
cally >99% as verified by SDS–PAGE. Selenomethionine-substituted
dShrm SD2 was expressed using PASM media (Studier, 2005), and
purification was essentially the same as for the native protein. Purifi-
cation of dRock SBD (724–938) was aided by the addition of an an-
ion exchange chromatography step before gel filtration.
Mutant mShrm3 and dShrm SD2 proteins
SC and HD mutations in mShrm3 and dShrm were made using the
QuikChange Site-Directed Mutagenesis Kit (Stratagene, Santa
Clara, CA). The mutant dShrm SD2 proteins were expressed and
purified in a manner similar to the wild type. All biochemical assays
with wild-type and HD proteins were performed with the indicated
protein fractions from gel filtration (Figure 3B). Gel filtration profiles
for Shrm SD2 proteins containing the SC1, SC2, or SC3 substitution
were are highly similar to that of wild-type SD2, with the exception
of some nucleic acid contamination in the SC1 and SC2 purifica-
tions. This was separated by gel filtration, and fractions correspond-
ing to the crystallized peak were used for all biochemical assays
(Supplementary Figure S5). For mShrm3 mutants, mutagenesis was
performed on mShrm3 in the pCS2-endolyn-Shrm3 expression plas-
mid. For in vitro expression of mShrm3 SD2 mutant proteins, the
mutated sequence encoding amino acids 1562–1986 was cloned
from the endolyn–Shrm3 vectors in pGex-2TK for expression in
E. coli CodonPlus (RIPL) cells. Recombinant proteins were expressed
and purified as described (Farber et al., 2011).
Crystallization of Drosophila Shroom SD2
Single thick, rod-shaped crystals were obtained for dShrm SD2 via
the vapor diffusion method with a reservoir solution containing
0.1 M 2-(N-morpholino)ethanesulfonic acid (MES) at pH 6.0, 1.35 M
K/Na tartrate, 0.7 M sodium thiocyanate, 11% glycerol (vol/vol), and
4 mM DTT. Crystals grew at 4°C in 7–10 d with a typical size of
80 × 40 × 500 μm and were cryoprotected by transition of the crystal
into a buffer containing 0.1 M MES, 1.4 M K/Na tartrate, 0.9 M so-
dium thiocynate, 15% glycerol, and 4 mM DTT. The cryoprotected
crystals were flash frozen under liquid nitrogen before data collec-
tion. The same procedure was used to crystallize and cryoprotect
SeMET-substituted SD2.
Structure determination
SD2 crystals belong to space group P2
1
2
1
2, with a = 72.6 Å, b =
85.6 Å, and c = 93.0 Å. Diffraction data from both native and SeMET
dShrm SD2 crystals were collected at beamline X25 at the National
Synchrotron Light Source, Brookhaven National Laboratory. Diffrac-
tion data integration, scaling, and merging were performed using
HKL2000 (Otwinowski and Minor, 1997). Initial phases were estimated
via the SAD method using SHELX C/D/E (Sheldrick, 2008), which
found six of the possible eight selenium sites. An initial model was
built into these experimental maps using Coot (Emsley and Cowtan,
2004). This model was then further refined against native data and
the model improved using a combination of simulated annealing and
positional, B factor, and TLS refinement (Zucker et al., 2010) within
Phenix (Adams et al., 2010). Model quality was monitored using Mol-
Probity (Davis et al., 2007). All structural images in this article were
generated using PyMOL (www.pymol.org). The coordinates and
structure factors for the Drosophila SD2 structure presented in this
Page 10
Volume 23 June 1, 2012 Structure of Shroom domain 2 | 2141
Cy3 or Cy5 maleimide (GE Healthcare) in cysteine labeling buffer
(20 mM HEPES, pH 7.6, 100 mM NaCl, 8% glycerol). Small (821–
938) dRock was labeled at C862 with Cy3 maleimide as described.
Large dRock (724–938) was labeled at the N-terminus with Cy5 suc-
cinimidyl ester (GE Healthcare) in amino labeling buffer. All labeling
reactions included 10× molar excess of fluorophore at room tem-
perature for 2 h. Excess fluorophore was removed from the samples
through extensive dialysis with labeling buffer. The labeling effi-
ciency was quantified using the extinction coefficient of the dye
compared with the protein concentration determined from a stan-
dard curve using a Bradford assay and found to be essentially 1:1.
FRET binding experiments
FRET titrations were performed in dShrm reaction buffer, using a
50 nM of Cy3-labeled dShrm or dRock and increasing concentra-
tions of Cy5-labeled dRock or dShrm. Cy3 was excited at 552 nM,
and the donor emission maximum (563 nm) was corrected for dilu-
tion, normalized, and plotted as a function of protein concentration
as the average of three independent experiments. Fluorescence
quenching (F
Q
) titrations were fitted to a single binding equation:
F
FdRock
dRock
Q
Q
d
=
×
+
[]
[]K
where F
Q
is the normalized change in donor fluorescence intensity
and K
d
is the dissociation constant.
ACKNOWLEDGMENTS
We thank Jeff Brodsky and Karen Arndt for critical comments on the
manuscript. Operations at the National Synchrotron Light Source
are supported by the Department of Energy, Office of Basic Energy
Research, and by the National Institutes of Health. Data collection at
the National Synchrotron Light Source was funded by the National
Center for Research Resources. This work was supported by Na-
tional Institutes of Health Grant GM097204.
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    • "When full length Shroom3 is expressed in COS7 cells, it binds actin filaments through a direct interaction with the SD1, resulting in co-localization with actin stress fibers (Fig. 3A) (Hildebrand, 2005 ). Consistent with previous results, coexpression of Shroom3 and the central coiled-coiled region of Rock results in the co-distribution of these proteins on actin stress fibers via the SD2 of Shroom3 and the Shroom-Binding Domain (SBD) of Rock (Fig. 3A) (Mohan et al., 2012; 2013). When Dvl2 and Shroom3 are co-expressed, Shroom3 is recruited to Dvl2 puncta (Fig. 3B ). "
    [Show abstract] [Hide abstract] ABSTRACT: Neural tube closure is a critical developmental event that relies on actomyosin contractility to facilitate specific processes such as apical constriction, tissue bending, and directional cell rearrangements. These complicated processes require the coordinated activities of Rho-Kinase (Rock), to regulate cytoskeletal dynamics and actomyosin contractility, and the Planar Cell Polarity (PCP) pathway, to direct the polarized cellular behaviors that drive convergent extension (CE) movements. Here we investigate the role of Shroom3 as a direct linker between PCP and actomyosin contractility during mouse neural tube morphogenesis. In embryos, simultaneous depletion of Shroom3 and the PCP components Vangl2 or Wnt5a results in an increased liability to NTDs and CE failure. We further show that these pathways intersect at Dishevelled, as Shroom3 and Dishevelled 2 co-distribute and form a physical complex in cells. We observed that multiple components of the Shroom3 pathway are planar polarized along mediolateral cell junctions in the neural plate of E8.5 embryos in a Shroom3 and PCP-dependent manner. Finally, we demonstrate that Shroom3 mutant embryos exhibit defects in planar cell arrangement during neural tube closure, suggesting a role for Shroom3 activity in CE. These findings support a model in which the Shroom3 and PCP pathways interact to control CE and polarized bending of the neural plate and provide a clear illustration of the complex genetic basis of NTDs. © 2015. Published by The Company of Biologists Ltd.
    Full-text · Article · Jan 2015 · Biology Open
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    • "Underlined amino acids constitute part of a conserved patch required for binding to Rock (Mohan et al., 2012). (E) Surface view of the Drosophila Shroom SD2 dimer as previous determined (Mohan et al., 2012) with the conserved arginine (R1474) residue in each monomer highlighted in green. "
    [Show abstract] [Hide abstract] ABSTRACT: Shroom3 is an actin-associated regulator of cell morphology that is required for neural tube closure, formation of the lens placode, and gut morphogenesis in mice and has been linked to chronic kidney disease and directional heart looping in humans. Numerous studies have shown that Shroom3 likely regulates these developmental processes by directly binding to Rho-kinase and facilitating the assembly of apically positioned contractile actomyosin networks. We have characterized the molecular basis for the neural tube defects caused by an ENU-induced mutation that results in an arginine-to-cysteine amino acid substitution at position 1838 of mouse Shroom3. We show that this substitution has no effect on Shroom3 expression or localization but ablates Rock binding and renders Shroom3 non-functional for the ability to regulate cell morphology. Our results indicate that Rock is the major downstream effector of Shroom3 in the process of neural tube morphogenesis. Based on sequence conservation and biochemical analysis, we predict that the Shroom-Rock interaction is highly conserved across animal evolution and represents a signaling module that is utilized in a variety of biological processes.
    Full-text · Article · Aug 2014 · Biology Open
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    • "Multiple lines of investigation, including biochemical, structural, and cellular analysis, have characterized and demonstrated the importance of the Shrm-Rock interaction in the regulation of cytoskeletal organization, cell shape, and tissue morphogenesis [5], [8]–[10], [37], [42]. Within Shrm, this interaction is mediated by a highly conserved Shrm-domain 2 (SD2) found at the C-termini of all Shrm proteins identified to date [35], [37], [42], [44]. "
    [Show abstract] [Hide abstract] ABSTRACT: Rho-associated coiled coil containing protein kinase (Rho-kinase or Rock) is a well-defined determinant of actin organization and dynamics in most animal cells characterized to date. One of the primary effectors of Rock is non-muscle myosin II. Activation of Rock results in increased contractility of myosin II and subsequent changes in actin architecture and cell morphology. The regulation of Rock is thought to occur via autoinhibition of the kinase domain via intramolecular interactions between the N-terminus and the C-terminus of the kinase. This autoinhibited state can be relieved via proteolytic cleavage, binding of lipids to a Pleckstrin Homology domain near the C-terminus, or binding of GTP-bound RhoA to the central coiled-coil region of Rock. Recent work has identified the Shroom family of proteins as an additional regulator of Rock either at the level of cellular distribution or catalytic activity or both. The Shroom-Rock complex is conserved in most animals and is essential for the formation of the neural tube, eye, and gut in vertebrates. To address the mechanism by which Shroom and Rock interact, we have solved the structure of the coiled-coil region of Rock that binds to Shroom proteins. Consistent with other observations, the Shroom binding domain is a parallel coiled-coil dimer. Using biochemical approaches, we have identified a large patch of residues that contribute to Shrm binding. Their orientation suggests that there may be two independent Shrm binding sites on opposing faces of the coiled-coil region of Rock. Finally, we show that the binding surface is essential for Rock colocalization with Shroom and for Shroom-mediated changes in cell morphology.
    Full-text · Article · Dec 2013 · PLoS ONE
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