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 contractility (Hildebrand, 2005
). Limited proteolysis of various SD2-containing protein fragments derived from mouse Shrm3 indicates the presence of a stable “core” of ~180 residues located at the C-terminus of SD2. We used these data to guide the design of SD2 expression constructs from several different Shrm proteins. We were able to obtain and optimize crystals from dShrm containing amino acid residues 1393–1576 () and determine its structure using the SAD method with selenomethionine (SeMET)-substituted crystals (see Material and Methods
and for a complete description of the structure determination procedure).
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 (more ...)
Data collection and refinement statistics for dShrm SD2.
The structure is refined at 2.7-Å resolution with an Rfree
value of 27.4%. The asymmetric unit contains a complete SD2 dimer, with 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 (). Each monomer contains three helices, 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 ( and Supplemental Figure S1). Within both the arm and body segments, coiled-coil interactions establish an extensive dimer interface, burying 4577 Å2
of surface area. This interface contains many conserved leucine and isoleucine residues, making interactions within the dimer interface reminiscent of leucine-zipper domains. In contrast to Shrm SD2, leucine 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 arrangements. To confirm that SD2 forms a dimer in solution, we treated purified SD2 with the chemical cross-linker glutaraldehyde and resolved the resulting species on SDS–PAGE (). These assays indicate that we can readily detect a dimeric SD2 species in solution (). 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 characterized SD2's solution state using gel filtration (). 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; and Supplemental 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 ( and Supplemental Figure S2). Structural homology searches failed to identify any structures 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 crystallization. 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 using 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), which is similar in length to the SD2s that are shown to cause apical constriction (Hildebrand, 2005
; Dietz et al., 2006
; ). 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 fragments are able to bind Rock, indicating that the crystallized fragment 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 (). 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 transmembrane 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 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 (). Therefore we conclude that the crystallized SD2 contains the Rock-binding site and, when properly localized, is sufficient to mediate apical constriction.
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 (more ...)
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 was formed by both SD2 chains. Given the large and extended dimerization interface, we were concerned that small perturbations, 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 binding. 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 identified two regions and generated multiple substitutions to target these regions ( and Supplemental Figure S1). We termed these variants homodimerization (HD) mutants. One interface mutant, HD1 (1468LLSL1471 to AASA; ), primarily targets the body segment, whereas the second HD mutant, HD2 (1546LIADARDL1553 to AAADARDA; ), primarily targets the coiled-coil within the arm segment. These amino acid changes are also predicted 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 secondary structure. We expressed and purified these proteins and compared their elution profile in gel filtration to wild-type protein (). We observe distinct changes with both mutants; protein containing the HD1 substitution elutes in a single broad peak distinct from both species observed 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 species. We isolated protein corresponding to dimer in the case of HD2 or to the majority peak from HD1 purification () and further characterized the effect of substitutions within the dimerization interface. We first tested their ability to form homodimers in solution by chemical cross-linking (). In this assay, both HD mutants exhibited 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. Consistent with the data obtained in the cross-linking experiment described 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 (). 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 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 composed of elements from both chains of the dimer.
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. (more ...)
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 (). 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 (). It should be noted that these patches are derived from amino acids residues on both the A and B chains, supporting the hypothesis that dimerization may be required to form a functional binding surface.
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 (more ...)
To address the importance of these surface clusters in Rock binding, we used the structural data to design amino acid substitutions within these potentially important surfaces. Given the preponderance of invariant residues, their broad distribution, and the 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 designed three multiresidue variants with alterations on the SD2 surface while avoiding residues that could play a role in dimerization. The surface cluster (SC) variants are 1402KMDEL1406 to AMDRA, 1470SLSERLA1476 to ALEEDLE, and 1509LKSDIERR1516 to AASDIEDA, which for clarity are named SC1, SC2, and SC3, respectively. The locations of these substitutions within the SD2 surface are indicated in (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 structure of the SD2 dimer (Supplemental Figure S5). We tested the surface cluster variants for their ability to bind dRock-SBD by pull-down (). 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 electrophoresis (). 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, indicating 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 dimer. Further, since the SC2 derivative exhibits an intermediate level of binding, we conclude that these amino acids may lie at the periphery 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 vertebrates. We noted that there was considerable sequence conservation 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: 1766KKAEL1770 to AKARA (SC1), 1834SLSGRLA1840 to ALEADLE (SC2), 1878LKENLDRR1885 to AAENLDDA (SC3), 1832LLSL1835 to AASA (HD1), and 1915LLIEQRKL1922 to ALIEQAKA (HD2). All of the homodimerization and surface cluster 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 (). In this assay, we could not detect binding of either of the homodimerization 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 ability of the surface cluster and homodimerization variants to form homodimers with an untagged, wild-type mShrm3-SD2 (). As expected from our studies with dShrm, the homodimerization mutations severely impaired dimerization, whereas the surface cluster mutations had no effect on binding to SD2. It should be noted that the surface cluster variant 1 bound with slightly reduced efficiency. On the basis of these data, we conclude that the Rock-binding 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 functional levels.
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 (more ...)
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 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 homodimerization 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 (, 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 pathway, 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 (). By measuring the increase in apical fluorescence relative to the decrease 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 surface cluster variants 2 or 3 caused apical constriction, and there was 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 apical 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 complex, 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 interaction (). Assuming a single-binding mode for this interaction, we calculate the equilibrium Kd
to be 0.58 ± 0.07 μM (). This affinity is comparable to that of RhoA, which has a reported Kd
of 0.13 μM (Blumenstein and Ahmadian, 2004
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 (more ...)
We next examined the stoichiometry of the dShrm–dRock complex. 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 (). Alternatively, complex was run on a gel filtration 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.