Chronophin coordinates cell leading edge dynamics by controlling active cofilin levels

Cofilin, a critical player of actin dynamics, is spatially and temporally regulated to control the direction and force of membrane extension required for cell locomotion. In carcinoma cells, although the signaling pathways regulating cofilin activity to control cell direction have been established, the molecular machinery required to generate the force of the protrusion remains unclear. We show that the cofilin phosphatase chronophin (CIN) spatiotemporally regulates cofilin activity at the cell edge to generate persistent membrane extension. We show that CIN translocates to the leading edge in a PI3-kinase–, Rac1-, and cofilin-dependent manner after EGF stimulation to activate cofilin, promotes actin free barbed end formation, accelerates actin turnover, and enhances membrane protrusion. In addition, we establish that CIN is crucial for the balance of protrusion/retraction events during cell migration. Thus, CIN coordinates the leading edge dynamics by controlling active cofilin levels to promote MTLn3 cell protrusion.Cofilin is one crucial mediator of actin cytoskeletal dynamics during cell motility (1–5). At the cell edge, cofilin severs F-actin filaments, generating substrates for Arp2/3-mediated branching activity and contributing to F-actin depolymerization by creating a new pointed end and F-actin assembly by increasing the pool of polymerization-competent actin monomers (G-actin) (6, 7). Because of its ability to sever actin filaments and thus, modulate actin dynamics, the precise spatial and temporal regulation of cofilin activity at the cell leading edge is crucial to cell protrusion, chemotaxis, and motility both in vitro and in vivo (2, 8–13). Misregulation of cofilin activity and/or expression is directly related to diseases, including tumor metastasis (14–18) and Alzheimer’s disease (19).Several mechanisms regulate tightly the activation of cofilin in response to upstream stimuli, including interaction with phosphatidylinositol (4,5)-bisphosphate (20–22), local pH changes (23, 24), and phosphorylation at a single regulatory serine (Ser3) (8, 25). The phosphorylation of cofilin, leading to its inactivation, is catalyzed by two kinase families: the LIM-kinases [LIMKs(Lin11, Isl-1, and Mec-3 domain)] and the testicular kinases (25–27). Two primary families of ser/thr phosphatases dephosphorylate and reactivate the actin-depolymerizing and -severing functions of cofilin: slingshot (SSH) (28) and chronophin (CIN) (29).SSH was identified as a cofilin phosphatase through genetic studies in Drosophila (28). The most active and abundant SSH isoform, SSH-1L, has been implicated in such biological processes as cell division, growth cone motility/morphology, neurite extension, and actin dynamics during membrane protrusion (30). SSH dephosphorylates a number of actin regulatory proteins in addition to cofilin, including LIMK1 (31) and Coronin 1B (32). CIN is a haloacid dehydrogenase-type phosphatase, a family of enzymes with activity in mammalian cells that has been poorly characterized. CIN dephosphorylates a very limited number of substrates (33) and as opposed to SSH, has little phosphatase activity toward LIMK both in vitro and in vivo; thus, it seems to be the more specific activator of cofilin (29, 30). CIN exhibits several predicted interaction motifs potentially linking it to regulation by PI3-kinase and phospholipase Cγ (PLCγ), both of which have been implicated in signaling to cofilin activation in vivo in MTLn3 adenocarcinoma cells (10, 34). CIN has been involved in cell division (29), cofilin–actin rod formation in neurons (35), and chemotaxing leukocytes (36, 37). The molecular mechanisms that control the activity and localization of CIN in cells are still not well-understood. In neutrophils, CIN mediates cofilin dephosphorylation downstream of Rac2 (36), and stimulation of protease-activated receptor2 results in recruitment of CIN and cofilin at the cell edge by β-arrestins to promote localized generation of free actin barbed ends, membrane protrusion, and chemotaxis (37). Chemotaxis to EGF by breast tumor cells is directly correlated with cancer cell invasion and metastasis (38, 39). Although cofilin activity is required for tumor cell migration, the contribution(s) of CIN to the regulation of actin dynamics at the leading edge has not yet been investigated.The importance of cofilin in regulating tumor cell motility has been extensively studied using MTLn3 mammary carcinoma cells as a model system. The initial step of MTLn3 cell chemotaxis to EGF consists of a biphasic actin polymerization response resulting from two peaks of free actin barbed end formation (34, 40, 41). The first or early peak of actin polymerization occurs at 1 min after EGF stimulation and requires both cofilin and PLCγ activities (34), but it is not dependent on cofilin dephosphorylation (42). This first transient allows the cells to sense EGF gradients and initiate small-membrane protrusions (11). The second or late peak of actin polymerization occurs at 3 min and is dependent on both cofilin and PI3-kinase activities (43, 44). Cofilin activity in this late transient has been associated with full protrusion of lamellipodia (34). The mechanism by which cofilin becomes activated at the 3-min peak has not been identified, although it is likely to involve the phosphoregulation of Ser3 (42, 45).In this work, we determine the molecular mechanisms involved in the full protrusion of the leading edge upon EGF stimulation. We have identified CIN as a critical regulator of cofilin activation to coordinate leading edge dynamics. Our results yield insights into how CIN controls cell protrusion, a key step in the process of cell migration and metastasis.     

Bimodal rheotactic behavior reflects flagellar beat asymmetry in human sperm cells

Rheotaxis, the directed response to fluid velocity gradients, has been shown to facilitate stable upstream swimming of mammalian sperm cells along solid surfaces, suggesting a robust physical mechanism for long-distance navigation during fertilization. However, the dynamics by which a human sperm orients itself relative to an ambient flow is poorly understood. Here, we combine microfluidic experiments with mathematical modeling and 3D flagellar beat reconstruction to quantify the response of individual sperm cells in time-varying flow fields. Single-cell tracking reveals two kinematically distinct swimming states that entail opposite turning behaviors under flow reversal. We constrain an effective 2D model for the turning dynamics through systematic large-scale parameter scans, and find good quantitative agreement with experiments at different shear rates and viscosities. Using a 3D reconstruction algorithm to identify the flagellar beat patterns causing left or right turning, we present comprehensive 3D data demonstrating the rolling dynamics of freely swimming sperm cells around their longitudinal axis. Contrary to current beliefs, this 3D analysis uncovers ambidextrous flagellar waveforms and shows that the cell’s turning direction is not defined by the rolling direction. Instead, the different rheotactic turning behaviors are linked to a broken mirror symmetry in the midpiece section, likely arising from a buckling instability. These results challenge current theoretical models of sperm locomotion.Taxis, the directed kinematic response to external signals, is a defining feature of living things that affects their reproduction, foraging, migration, and survival strategies (1–4). Higher organisms rely on sophisticated networks of finely tuned sensory mechanisms to move efficiently in the presence of chemical or physical stimuli. However, various fundamental forms of taxis are already manifest at the unicellular level, ranging from chemotaxis in bacteria (5) and phototaxis in unicellular green algae (2) to the mechanical response (durotaxis) of fibroblasts (6) and rheotaxis (7, 8) in spermatozoa (3, 9–12). Over the last few decades, much progress has been made in deciphering chemotactic, phototactic, and durotactic pathways in prokaryotic and eukaryotic model systems. In contrast, comparatively little is known about the physical mechanisms that enable flow gradient sensing in sperm cells (3, 9–13). Recent studies (3, 12) suggest that mammalian sperm use rheotaxis for long-distance navigation, but it remains unclear how shear flows alter flagellar beat patterns in the vicinity of surfaces and, in particular, how such changes in the beat dynamics affect the steering process. Answering these questions will be essential for evaluating the importance of chemical (14) and physical (4) signals during mammalian fertilization (15–17).A necessary requirement for any form of directed kinematic response is the ability to change the direction of locomotion. Multiflagellate bacteria achieve this feat by varying their motor activity, resulting in alternating phases of entangled and disentangled flagellar dynamics that give rise to run-and-tumble motion (5). A similar mechanism was recently discovered in the biflagellate eukaryote Chlamydomonas reinhardtii (18). This unicellular green alga actively redirects its swimming motion through occasional desynchronization of its two cilia (19), although it is still debated whether this effect is of mechanical (20) or hydrodynamic (21, 22) origin. Experiments (23) show that the alga’s reorientation dynamics can lead to localization in shear flow (24, 25), with potentially profound implications in marine ecology. In contrast to taxis in multiflagellate organisms (2, 5, 18, 26, 27), the navigation strategies of uniflagellate cells are less well understood. For instance, it was discovered only recently that uniflagellate marine bacteria, such as Vibrio alginolyticus and Pseudoalteromonas haloplanktis, use a buckling instability in their lone flagellum to change their swimming direction (28). However, as passive prokaryotic flagella differ fundamentally from their active eukaryotic counterparts, it is unclear to what extent such insights translate to spermatozoa.Earlier studies of human sperm locomotion have identified several potential steering and transport mechanisms, including thermotaxis (4), uterine peristalsis (29, 30), and chemotaxis (14, 16, 31), but their relative importance has yet to be quantified. Recent experiments (3, 32, 33) demonstrate that rheotaxis, combined with steric surface alignment (12, 34), enables robust long-distance navigation by turning sperm cells preferentially against an externally imposed flow direction (9, 10), but how exactly this realignment process happens is unknown. It has been suggested (32, 35, 36) that the intrinsic curvature or chiral beat dynamics (37, 38) of the flagellum could play an essential role in rheotactic steering, but this remains to be confirmed in experiments. Similarly, an increasing number of theoretical models (36, 39–47) still await empirical validation, because 3D data for the beat pattern of sperm swimming close to surfaces has been lacking.To examine the dynamics of human sperm rheotaxis quantitatively, we here combine microfluidic experiments with mathematical modeling and 3D flagellar beat reconstruction. Single-cell tracking reveals the existence of two kinematically distinct swimming states that result in opposite turning behaviors under flow reversal. We quantify this effect for a range of viscosities and shear rates, and use these comprehensive data to constrain an effective 2D model through a systematic large-scale scan ( > 6,000 parameter combinations). To identify the details of the flagellar beat dynamics during rheotaxis, we developed an algorithm that translates 2D intensity profiles into 3D positional data. Our 3D analysis confirms that human sperm perform a rolling motion (48), characterized by weakly nonplanar beat patterns and a rotating beat plane. However, contrary to current beliefs, we find that neither the rolling direction nor beat helicity determine the turning direction after flow reversal. Instead, the rheotactic turning behavior correlates with a previously unrecognized asymmetry in the midpiece, likely caused by a buckling instability. These findings call for a revision and extension of current models (36, 39–44, 46).     

Conformational dynamics of a crystalline protein from microsecond-scale molecular dynamics simulations and diffuse X-ray scattering

X-ray diffraction from protein crystals includes both sharply peaked Bragg reflections and diffuse intensity between the peaks. The information in Bragg scattering is limited to what is available in the mean electron density. The diffuse scattering arises from correlations in the electron density variations and therefore contains information about collective motions in proteins. Previous studies using molecular-dynamics (MD) simulations to model diffuse scattering have been hindered by insufficient sampling of the conformational ensemble. To overcome this issue, we have performed a 1.1-μs MD simulation of crystalline staphylococcal nuclease, providing 100-fold more sampling than previous studies. This simulation enables reproducible calculations of the diffuse intensity and predicts functionally important motions, including transitions among at least eight metastable states with different active-site geometries. The total diffuse intensity calculated using the MD model is highly correlated with the experimental data. In particular, there is excellent agreement for the isotropic component of the diffuse intensity, and substantial but weaker agreement for the anisotropic component. Decomposition of the MD model into protein and solvent components indicates that protein–solvent interactions contribute substantially to the overall diffuse intensity. We conclude that diffuse scattering can be used to validate predictions from MD simulations and can provide information to improve MD models of protein motions.Proteins explore many conformations while carrying out their functions in biological systems (1–3). X-ray crystallography is the dominant source of information about protein structure; however, crystal structure models usually consist of just a single major conformation and at most a small portion of the model as alternate conformations. Crystal structures therefore are missing many details about the underlying conformational ensemble (4).Proteins assembled in crystalline arrays, like proteins in solution, exhibit rich conformational diversity (4) and often can perform their native functions (5). Many methods have emerged for using Bragg data to model conformational diversity in protein crystals (6–17). The development of these methods has been important as conformational diversity can lead to inaccuracies in protein structure models (9, 18–20). A key limitation of using the Bragg data, however, is that different models of conformational diversity can yield the same mean electron density.Whereas the Bragg scattering only contains information about the mean electron density, diffuse scattering (diffraction resulting in intensity between the Bragg peaks) is sensitive to spatial correlations in electron density variations (21–28) and therefore contains information about the way that atomic positions vary together in protein crystals. Because models that yield the same mean electron density can yield different correlations in electron density variations, diffuse scattering provides a means to increase the accuracy of crystallography for determining protein conformational variations (29). Peter Moore (30) and Mark Wilson (31) have argued that diffuse scattering should be used to test models of conformational diversity in X-ray crystallography.Several pioneering studies used diffuse scattering to reveal insights into correlated motions in proteins (17, 30, 32–49). Some of these studies used diffuse scattering to experimentally validate predictions of correlated motions from molecular-dynamics (MD) simulations (35–37, 40, 42–44). These studies revealed important insights but were limited by inadequate sampling of the conformational ensemble, leading to lack of convergence of the diffuse scattering calculations (35). Microsecond-scale simulations of staphylococcal nuclease were predicted to be adequate for convergence of diffuse scattering calculations (42). Modern simulation algorithms and computer hardware now enable microsecond or longer MD simulations of protein crystals (50).Here, we present calculations of diffuse X-ray scattering using a 1.1-μs MD simulation of crystalline staphylococcal nuclease. The results demonstrate that we have overcome the past limitation of inadequate sampling. We chose staphylococcal nuclease because the experiments of Wall et al. (49) still represent the only complete, high-quality, 3D diffuse scattering data set from a protein crystal. The calculated diffuse intensity is very similar using two independent halves of the trajectory; the results therefore are reproducible and can be meaningfully compared with the experimental data. The MD simulation provides a rich picture of conformational diversity in the energy landscape of a protein crystal, consisting of at least eight metastable states. Like previous MD studies of crystalline staphylococcal nuclease (42–44), the agreement of the simulation with the total experimental diffuse intensity is excellent, supporting the use of MD simulations to model diffuse scattering data. Unlike previous MD studies, we separately compared the more finely structured, anisotropic component of the diffuse intensity with experimental data. The agreement is substantial but weaker than for the isotropic component, indicating there are inaccuracies in the MD models. Our results therefore point toward using diffuse scattering to improve MD models of protein motions.     

Nanomechanical mechanism for lipid bilayer damage induced by carbon nanotubes confined in intracellular vesicles

Understanding the behavior of low-dimensional nanomaterials confined in intracellular vesicles has been limited by the resolution of bioimaging techniques and the complex nature of the problem. Recent studies report that long, stiff carbon nanotubes are more cytotoxic than flexible varieties, but the mechanistic link between stiffness and cytotoxicity is not understood. Here we combine analytical modeling, molecular dynamics simulations, and in vitro intracellular imaging methods to reveal 1D carbon nanotube behavior within intracellular vesicles. We show that stiff nanotubes beyond a critical length are compressed by lysosomal membranes causing persistent tip contact with the inner membrane leaflet, leading to lipid extraction, lysosomal permeabilization, release of cathepsin B (a lysosomal protease) into the cytoplasm, and cell death. The precise material parameters needed to activate this unique mechanical pathway of nanomaterials interaction with intracellular vesicles were identified through coupled modeling, simulation, and experimental studies on carbon nanomaterials with wide variation in size, shape, and stiffness, leading to a generalized classification diagram for 1D nanocarbons that distinguishes pathogenic from biocompatible varieties based on a nanomechanical buckling criterion. For a wide variety of other 1D material classes (metal, oxide, polymer), this generalized classification diagram shows a critical threshold in length/width space that represents a transition from biologically soft to stiff, and thus identifies the important subset of all 1D materials with the potential to induce lysosomal permeability by the nanomechanical mechanism under investigation.The interactions of low-dimensional materials with the external or plasma membrane of living cells have been the subject of prior studies due to their importance in uptake and delivery, antibacterial action, and nanomaterial safety (1–6). Following uptake, nanomaterials may also interact with internal membranes while under confinement in intracellular vesicles (7–10), but the biophysics of these geometrically constrained systems is poorly understood. Low-dimensional materials interact with biological systems in complex ways dictated by their 1D nanofibrous or 2D nanosheet geometries (7, 11–20). These interactions typically begin when materials encounter the plasma membrane and initiate phenomena that can include adhesion, membrane deformation, penetration, lipid extraction, entry, frustrated uptake, or cytotoxicity (4, 11–14, 19–21). Recent experimental data suggest that the cellular response to some 1D materials is governed by their interaction with the internal lipid-bilayer membranes of endosomes and lysosomes following nanomaterial uptake (7–10). The resulting geometry is fundamentally different in that the fibrous materials are confined within a vesicle, imposing geometric constraints and introducing mechanical forces that act bidirectionally––i.e., on both the thin fibrous structure and the inner leaflet of the soft membrane. The fundamental biophysics of this tube-in-vesicle system is virtually unexplored, yet may be critical for understanding the cellular response to nanotubes/fibers, where shape and stiffness are among the known determinants of toxicity (13, 21). The technique of coarse-grained molecular dynamics (MD), demonstrated to be effective in the study of complex biomolecular systems (22, 23), has been applied to whole lipid-bilayer patches to reveal a biophysical mechanism for carbon nanotube interaction with the plasma membranes leading to tip entry and uptake (4, 19, 20). The same technique may also provide insight relevant to internal membrane interactions, although whole vesicle MD is a significant challenge. Here we use a complement of techniques including coarse-grained MD, all-atom MD, in vitro bioimaging, and carbon nanotube length modification to reveal the behavior of vesicle-encapsulated carbon nanotubes and identify the conditions and carbon nanotube (CNT) types that lead to mechanical stress and membrane damage following cellular uptake and packaging in lysosomes (8).     

Understanding the kinetic mechanism of RNA single base pair formation

RNA functions are intrinsically tied to folding kinetics. The most elementary step in RNA folding is the closing and opening of a base pair. Understanding this elementary rate process is the basis for RNA folding kinetics studies. Previous studies mostly focused on the unfolding of base pairs. Here, based on a hybrid approach, we investigate the folding process at level of single base pairing/stacking. The study, which integrates molecular dynamics simulation, kinetic Monte Carlo simulation, and master equation methods, uncovers two alternative dominant pathways: Starting from the unfolded state, the nucleotide backbone first folds to the native conformation, followed by subsequent adjustment of the base conformation. During the base conformational rearrangement, the backbone either retains the native conformation or switches to nonnative conformations in order to lower the kinetic barrier for base rearrangement. The method enables quantification of kinetic partitioning among the different pathways. Moreover, the simulation reveals several intriguing ion binding/dissociation signatures for the conformational changes. Our approach may be useful for developing a base pair opening/closing rate model.RNAs perform critical cellular functions at the level of gene expression and regulation (1–4). RNA functions are determined not only by RNA structure or structure motifs [e.g., tetraloop hairpins (5, 6)] but also by conformational distributions and dynamics and kinetics of conformational changes. For example, riboswitches can adopt different conformations in response to specific conditions of the cellular environment (7, 8). Understanding the kinetics, such as the rate and pathways for the conformational changes, is critical for deciphering the mechanism of RNA function (9–19). Extensive experimental and theoretical studies on RNA folding kinetics have provided significant insights into the kinetic mechanism of RNA functions (19–36). However, due to the complexity of the RNA folding energy landscape (37–46) and the limitations of experimental tools (47–55), many fundamental problems, including single base flipping and base pair formation and fraying, remain unresolved. These unsolved fundamental problems have hampered our ability to resolve other important issues, such as RNA hairpin and larger structure folding kinetics. Several key questions remain unanswered, such as whether the hairpin folding is rate-limited by the conformational search of the native base pairs, whose formation leads to fast downhill folding of the whole structure, or by the breaking of misfolded base pairs before refolding to the native structure (18, 19, 54–73).Motivated by the need to understand the basic steps of nucleic acids folding, Hagan et al. (74) performed forty-three 200-ps unfolding trajectories at 400 K and identified both on- and off-pathway intermediates and two dominant unfolding pathways for a terminal C-G base pair in a DNA duplex. In one of the pathways, base pairing and stacking interactions are broken concomitantly, whereas in the other pathway, base stacking is broken after base pairing is disrupted. Furthermore, the unfolding requires that the Cyt diffuse away from the pairing Gua to a distance such that the C-G hydrogen bond cannot reform easily. More recently, Colizzi and Bussi (75) performed molecular dynamics (MD) pulling simulations for an RNA duplex and construct free energy landscape from the pulling simulation. The simulation showed that the base pair opening reaction starts with the unbinding of the 5′-base, followed by the unbinding of the 3′-base (i.e., the 5′-base is less stable than the 3′-base). These previous unfolding simulations offered significant insights into the pathways and transition states. However, as shown below, several important issues remain.One intriguing problem is the rate model for base pairing. There are currently three main types of models. In the first type of model, the barrier ΔG+‡ for closing a base pair is dominated by the entropic cost ΔS for positioning the nucleotides to the base-paired configuration and the barrier ΔG−‡ for opening a base pair is the enthalpic cost ΔH for disrupting the hydrogen bonds and base stacking interactions (18, 59, 60). In the second type of model, ΔG+‡ is the net free energy change for base pairing ΔG = ΔH ? TΔS and ΔG−‡ is zero (76, 77). In the third type of model, ΔG±‡=±ΔG/2 is used (78). In addition to the above three main types, other models, such as more sophisticated hybrid rate models, have been proposed (29).In this paper, we report a hybrid method (see Fig. 1) to investigate the single base pairing process. In contrast to the previous simulations for temperature- or force-induced unfolding reactions, we directly model the folding process here (i.e., the base pair closing process). Specifically, we use MD simulations to identify the conformational clusters. Based on the network of the conformational clusters as a reduced conformational ensemble, we apply kinetic Monte Carlo (KMC) and master equation (ME) methods to elucidate the detailed roles of base pairing and stacking interactions, as well as the roles of water and ions (79–82). The study reveals previously unidentified kinetics pathways, misfolded states, and rate-limiting steps. A clear understanding of the microscopic details of the elementary kinetic move is a prerequisite for further rigorous study of large-scale RNA kinetic studies. The method described here may provide a feasible way to develop a rate model for the base pair/stack-based kinetic move set. Furthermore, the mechanism of RNA single base folding may provide useful insights into many biologically significant processes, such as nucleotide flipping (83) in helicases and base pair fraying (84) (as the possible first step for nucleic duplex melting in nucleic acid enzymatic processes).Open in a separate windowFig. 1.(A) Folding of a single nucleotide (G1, red) from the unfolded (Left) to the native folded (Right) state. (B) Exhaustive sampling for the (discrete) conformations of the G1 nucleotide (Right) through enumeration of the torsion angles (formed by the blue bonds). (C) Schematic plot shows the trajectories on the energy landscape (depicted with two reaction coordinates for clarity) explored by the MD simulations. The lines, open circles, and hexagons denote the trajectories; the initial states; and the (centroid structures of the) clusters, respectively. (D) Conformational network based on six clusters. (E) The rmsds to the different clusters provide information about the structural changes in a MD trajectory.     

Visualization of two transfer RNAs trapped in transit during elongation factor G-mediated translocation

During protein synthesis, coupled translocation of messenger RNAs (mRNA) and transfer RNAs (tRNA) through the ribosome takes place following formation of each peptide bond. The reaction is facilitated by large-scale conformational changes within the ribosomal complex and catalyzed by elongtion factor G (EF-G). Previous structural analysis of the interaction of EF-G with the ribosome used either model complexes containing no tRNA or only a single tRNA, or complexes where EF-G was directly bound to ribosomes in the posttranslocational state. Here, we present a multiparticle cryo-EM reconstruction of a translocation intermediate containing two tRNAs trapped in transit, bound in chimeric intrasubunit ap/P and pe/E hybrid states. The downstream ap/P-tRNA is contacted by domain IV of EF-G and P-site elements within the 30S subunit body, whereas the upstream pe/E-tRNA maintains tight interactions with P-site elements of the swiveled 30S head. Remarkably, a tight compaction of the tRNA pair can be seen in this state. The translocational intermediate presented here represents a previously missing link in understanding the mechanism of translocation, revealing that the ribosome uses two distinct molecular ratchets, involving both intra- and intersubunit rotational movements, to drive the synchronous movement of tRNAs and mRNA.During protein synthesis the ribosome iteratively incorporates new amino acids delivered by aminoacylated transfer RNAs (tRNA) into the growing polypeptide chain in a manner specified by the codons in a messenger RNAs (mRNA). This elongation cycle is controlled by the two translocational GTPases elongation factors (EF)-Tu and EF-G. Following EF-Tu–dependent delivery of aminoacyl-tRNA to the A site and peptide bond formation, the ribosome adopts a pretranslocational state containing a peptidyl A-site tRNA and a deacylated P-site tRNA. In the subsequent translocation reaction, the interplay between the ribosome and elongation factor EF-G shifts the tRNAs from the A and P sites to the P and E sites, respectively. In each of these binding sites a tRNA contacts both ribosomal subunits and interacts with the 30S and 50S subunits via its anticodon-stem loop (ASL) and acceptor arm, respectively (1). Partial tRNA movement can occur before the EF-G–dependent translocation step, involving spontaneous and reversible movement of the tRNA acceptor arms relative to the large ribosomal subunit, which leads to a shift from classic A/A and P/P binding states into intersubunit A/P and P/E hybrid states (where the first and second letters indicate tRNA contacts on the small and large subunits, respectively) (2–4).A remarkable feature of translocation is the precise coupling of movement of the tRNAs together with the bound mRNA (designated as the tRNA₂•mRNA module), so that the mRNA advances by exactly one codon on the ribosome. Translocation is associated with large-scale conformational changes within the ribosomal complex, which includes rotation and back-rotation between the two subunits (during which the small subunit rotates counterclockwise and clockwise relative to the large subunit, viewing the ribosome from the solvent side of the small subunit) (5–7), and an additional forward and reverse swiveling movement of the 30S head (an autonomous domain of the 30S subunit, which can rotate around an axis roughly orthogonal to the axis of intersubunit rotation) (6, 8–13). Structural studies have suggested that intersubunit rotation within the pretranslocational complex is coupled to tRNA hybrid state formation (14–18). EF-G–dependent movement of the tRNAs and mRNA on the 30S subunit then occurs during reversal of the intersubunit rotation (6, 7). Moreover, multiparticle cryo-EM and X-ray crystallography studies suggest that movement of the tRNAs relative to the 30S subunit occurs via additional intermediate tRNA binding states, which are formed upon the back rotation of the 30S-body/platform and a large swiveling movement of the 30S head (6, 10). One of the first implications of swiveling of the 30S subunit head came from the studies of Schuwirth et al. (19), who observed a constriction of 13 Å between head and body of the 30S subunit that would block passage of tRNA between the P and E sites. These authors suggested that rotation of the 30S head would allow movement of the tRNA ASL, and could correspond to an unlocking event during translocation. Although swiveling of the small subunit head has been observed in different ribosomal complexes with bound EF-G (or eEF2) (6, 8–13) or with bound tRNAs (18), it has not been observed directly in the context of an authentic translocation complex containing EF-G together with two tRNAs. Previous structural analysis of translocation used model complexes where EF-G was directly bound to either vacant ribosomes, to ribosomal complexes with one tRNA, or to complexes in the posttranslocational state (5, 6, 8, 10–13, 20–22). The present study describes a cryo-EM reconstruction more closely resembling an authentic translocation intermediate, in which EF-G•GTP was bound to a canonical pretranslocational ribosomal complex containing two tRNAs and mRNA, and stalled during the translocation reaction by the antibiotic fusidic acid (FA). The resulting sample was analyzed by means of multiparticle cryo-EM (23). The classification yielded only a single major population of 70S•EF-G•GDP•FA particles trapped in an intermediate state of the translocation reaction. In contrast to all previously described 70S•EF-G structures (5, 6, 10–13, 20–22), the resulting reconstruction directly visualizes two tRNAs bound to the ribosome in two different chimeric intrasubunit hybrid states. The data presented here show how reciprocal conformational changes within the ribosome coordinate the synchronous movement of the mRNA and bound tRNA pair.     

Remodeling nuclear architecture allows efficient transport of herpesvirus capsids by diffusion

The nuclear chromatin structure confines the movement of large macromolecular complexes to interchromatin corrals. Herpesvirus capsids of approximately 125 nm assemble in the nucleoplasm and must reach the nuclear membranes for egress. Previous studies concluded that nuclear herpesvirus capsid motility is active, directed, and based on nuclear filamentous actin, suggesting that large nuclear complexes need metabolic energy to escape nuclear entrapment. However, this hypothesis has recently been challenged. Commonly used microscopy techniques do not allow the imaging of rapid nuclear particle motility with sufficient spatiotemporal resolution. Here, we use a rotating, oblique light sheet, which we dubbed a ring-sheet, to image and track viral capsids with high temporal and spatial resolution. We do not find any evidence for directed transport. Instead, infection with different herpesviruses induced an enlargement of interchromatin domains and allowed particles to diffuse unrestricted over longer distances, thereby facilitating nuclear egress for a larger fraction of capsids.The nucleus is structured into chromatin and interchromatin compartments, giving it a “sponge-like” appearance, with the chromatin representing the actual sponge material and the interchromatin regions representing the enclosed pores, tubes, tunnels, and cisterna. This conceptual model has important implications for how differently sized molecules could move inside the nucleus. Although small molecules like GFP, streptavidin, fluorescent dextrans, or mRNA can mostly freely diffuse throughout the nucleus (1–3), large macromolecular assemblies, such as 100-nm fluorescent beads and promyelocytic leukemia (PML) or Cajal bodies, are trapped in interchromatin spaces called corrals. These corrals slowly move as a result of chromatin dynamics, and particles can also escape over long time scales (4, 5). Herpesvirus capsids of 125-nm diameter are assembled in the nucleus and must move to the nuclear periphery to exit the nucleus by budding through the nuclear membranes (6). Similarly, large host cargo like ribonucleoprotein (RNP) particles may need to move through the nucleus before they exit the nucleus in the same way (7). Earlier, we and others provided data that suggested that herpesvirus capsids use an active, directed mechanism based on F-actin to transport through the nucleoplasm (8, 9) and/or that nuclear F-actin is involved in capsid assembly. However, we recently showed that herpesviruses do not induce nuclear F-actin in most cells, and, more importantly, that nuclear capsid motility is not dependent on nuclear F-actin (10). This led us to reinvestigate nuclear capsid motility by single particle tracking to address whether molecular motors power it.During herpesvirus infection, the interchromatin domains enlarge and the nuclear volume increases as much as twofold (11). At the same time, viral replication compartments expand, move, and coalesce, but do not mix (12–16). By using a light-sheet modality we dubbed a ring-sheet, we were able to trace intranuclear herpes virus capsids with high spatial and temporal precision and low cytotoxicity. Our data show that nuclear herpesvirus capsids do not use a directed transport mechanism. Instead, we find by single particle tracking and analysis of capsid motility that capsids diffuse in enlarged nuclear spaces. We conclude that the previously described size increase of the interchromatin domain increases the likelihood of capsids to reach the nuclear envelope by diffusion.     

Efficient plant male fertility depends on vegetative nuclear movement mediated by two families of plant outer nuclear membrane proteins

Increasing evidence suggests that nuclear migration is important for eukaryotic development. Although nuclear migration is conserved in plants, its importance for plant development has not yet been established. The most extraordinary plant nuclear migration events involve plant fertilization, which is starkly different from that of animals. Instead of evolving self-propelled sperm cells (SCs), plants use pollen tubes to deliver SCs, in which the pollen vegetative nucleus (VN) and the SCs migrate as a unit toward the ovules, a fundamental but barely understood process. Here, we report that WPP domain-interacting proteins (WIPs) and their binding partners the WPP domain-interacting tail-anchored proteins (WITs) are essential for pollen nuclear migration. Loss-of-function mutations in WIT and/or WIP gene families resulted in impaired VN movement, inefficient SC delivery, and defects in pollen tube reception. WIPs are Klarsicht/ANC-1/Syne-1 Homology (KASH) analogs in plants. KASH proteins are key players in animal nuclear migration. Thus, this study not only reveals an important nuclear migration mechanism in plant fertilization but also, suggests that similar nuclear migration machinery is conserved between plants and animals.Nuclear migration is essential for cell differentiation, polarization, and migration, which influence organism development (1–3). Examples range from Caenorhabditis elegans P-cell development to mammalian neural development (1–3). The key players in opisthokont nuclear migration are the inner nuclear membrane Sad1/UNC-84 (SUN) proteins and outer nuclear membrane Klarsicht/ANC-1/Syne-1 Homology (KASH) proteins. SUN and KASH proteins form the linkers of the nucleoskeleton and the cytoskeleton complexes at the nuclear envelope (NE) and transfer cytoplasmic forces to the nucleus (1–3). In plants, nuclear migration is associated with a number of developmental events and environmental responses, including fertilization, root and leaf hair formation, and plant–microbe interactions (4, 5). So far, little is known about the mechanism of plant nuclear migration. Although SUN proteins are conserved in plants (6, 7), absence of animal KASH homologs in plants suggests that plants may have evolved different molecular solutions to achieve nuclear migration. Recently, WPP domain-interacting proteins (WIPs) were identified as KASH proteins in plants (8), and their outer nuclear membrane binding partners WPP domain-interacting tail anchored proteins (WITs) were shown to interact with myosin XI-I (9). The WIT–myosin XI-I complexes regulate nuclear movement in root and mesophyll cells, but no developmental events have been linked to these nuclear movements (9).Essential for plant fertility, pollen tube growth harbors the most dramatic nuclear movement in plants. Unlike animals, which have sperm cells (SCs) that travel through self-propelled flagellum, flowering plants use pollen tubes to deliver SCs to ovules (10–13). In Arabidopsis, pollen tube growth is guided by chemical cues in carpel tissues and attracted by small peptides secreted by synergid cells in the vicinity of ovules (14–18). Pollen tube reception is completed by pollen tube burst, SC release, and degeneration of synergid cells (12). If this process fails, a second pollen tube can be attracted to the same ovule for a second attempt, resulting in polytubey (19). The SCs [or their progenitor the generative cell (GC)] are enclosed by an endocytic membrane tethered to the pollen vegetative nucleus (VN) (20). During pollen tube elongation, the VN and the SCs/GC are usually closely associated and move as a male germ unit (MGU) (13, 21). For decades, the movement of the MGU has been analyzed using cytoskeleton-depolymerizing reagents or heterogeneous antimyosin antibodies (22–27). However, no genes have been implicated in MGU movement, and the function of the joint migration of VN and GC/SC remains hypothetical.Here, we have identified the Arabidopsis WIT and WIP protein families as key players in VN movement. WIP1 and WIT1 are localized at the vegetative nuclear envelope (VNE). Loss of either WIT or WIP family proteins impaired VN movement, resulting in defective pollen tube reception and inefficient SC-to-ovule migration. This study has not only identified a molecular mechanism regulating the VN movement but also, revealed an important function of the VN in plant fertilization.     

Self-assembly of alkynylplatinum(II) terpyridine amphiphiles into nanostructures via steric control and metal–metal interactions

A series of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing the hydrophilic oligo(para-phenylene ethynylene) with two 3,6,9-trioxadec-1-yloxy chains was designed and synthesized. The mononuclear alkynylplatinum(II) terpyridine complex was found to display a very strong tendency toward the formation of supramolecular structures. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would lead to the formation of nanotubes or helical ribbons. These desirable nanostructures were found to be governed by the steric bulk on the platinum(II) terpyridine moieties, which modulates the directional metal−metal interactions and controls the formation of nanotubes or helical ribbons. Detailed analysis of temperature-dependent UV-visible absorption spectra of the nanostructured tubular aggregates also provided insights into the assembly mechanism and showed the role of metal−metal interactions in the cooperative supramolecular polymerization of the amphiphilic platinum(II) complexes.Square-planar d⁸ platinum(II) polypyridine complexes have long been known to exhibit intriguing spectroscopic and luminescence properties (1–54) as well as interesting solid-state polymorphism associated with metal−metal and π−π stacking interactions (1–14, 25). Earlier work by our group showed the first example, to our knowledge, of an alkynylplatinum(II) terpyridine system [Pt(tpy)(C ≡ CR)]⁺ that incorporates σ-donating and solubilizing alkynyl ligands together with the formation of Pt···Pt interactions to exhibit notable color changes and luminescence enhancements on solvent composition change (25) and polyelectrolyte addition (26). This approach has provided access to the alkynylplatinum(II) terpyridine and other related cyclometalated platinum(II) complexes, with functionalities that can self-assemble into metallogels (27–31), liquid crystals (32, 33), and other different molecular architectures, such as hairpin conformation (34), helices (35–38), nanostructures (39–45), and molecular tweezers (46, 47), as well as having a wide range of applications in molecular recognition (48–52), biomolecular labeling (48–52), and materials science (53, 54). Recently, metal-containing amphiphiles have also emerged as a building block for supramolecular architectures (42–44, 55–59). Their self-assembly has always been found to yield different molecular architectures with unprecedented complexity through the multiple noncovalent interactions on the introduction of external stimuli (42–44, 55–59).Helical architecture is one of the most exciting self-assembled morphologies because of the uniqueness for the functional and topological properties (60–69). Helical ribbons composed of amphiphiles, such as diacetylenic lipids, glutamates, and peptide-based amphiphiles, are often precursors for the growth of tubular structures on an increase in the width or the merging of the edges of ribbons (64, 65). Recently, the optimization of nanotube formation vs. helical nanostructures has aroused considerable interests and can be achieved through a fine interplay of the influence on the amphiphilic property of molecules (66), choice of counteranions (67, 68), or pH values of the media (69), which would govern the self-assembly of molecules into desirable aggregates of helical ribbons or nanotube scaffolds. However, a precise control of supramolecular morphology between helical ribbons and nanotubes remains challenging, particularly for the polycyclic aromatics in the field of molecular assembly (64–69). Oligo(para-phenylene ethynylene)s (OPEs) with solely π−π stacking interactions are well-recognized to self-assemble into supramolecular system of various nanostructures but rarely result in the formation of tubular scaffolds (70–73). In view of the rich photophysical properties of square-planar d⁸ platinum(II) systems and their propensity toward formation of directional Pt···Pt interactions in distinctive morphologies (27–31, 39–45), it is anticipated that such directional and noncovalent metal−metal interactions might be capable of directing or dictating molecular ordering and alignment to give desirable nanostructures of helical ribbons or nanotubes in a precise and controllable manner.Herein, we report the design and synthesis of mono- and dinuclear alkynylplatinum(II) terpyridine complexes containing hydrophilic OPEs with two 3,6,9-trioxadec-1-yloxy chains. The mononuclear alkynylplatinum(II) terpyridine complex with amphiphilic property is found to show a strong tendency toward the formation of supramolecular structures on diffusion of diethyl ether in dichloromethane or dimethyl sulfoxide (DMSO) solution. Interestingly, additional end-capping with another platinum(II) terpyridine moiety of various steric bulk at the terminal alkyne would result in nanotubes or helical ribbons in the self-assembly process. To the best of our knowledge, this finding represents the first example of the utilization of the steric bulk of the moieties, which modulates the formation of directional metal−metal interactions to precisely control the formation of nanotubes or helical ribbons in the self-assembly process. Application of the nucleation–elongation model into this assembly process by UV-visible (UV-vis) absorption spectroscopic studies has elucidated the nature of the molecular self-assembly, and more importantly, it has revealed the role of metal−metal interactions in the formation of these two types of nanostructures.     

Structural insights into the histone H1-nucleosome complex

Linker H1 histones facilitate formation of higher-order chromatin structures and play important roles in various cell functions. Despite several decades of effort, the structural basis of how H1 interacts with the nucleosome remains elusive. Here, we investigated Drosophila H1 in complex with the nucleosome, using solution nuclear magnetic resonance spectroscopy and other biophysical methods. We found that the globular domain of H1 bridges the nucleosome core and one 10-base pair linker DNA asymmetrically, with its α3 helix facing the nucleosomal DNA near the dyad axis. Two short regions in the C-terminal tail of H1 and the C-terminal tail of one of the two H2A histones are also involved in the formation of the H1–nucleosome complex. Our results lead to a residue-specific structural model for the globular domain of the Drosophila H1 in complex with the nucleosome, which is different from all previous experiment-based models and has implications for chromatin dynamics in vivo.Eukaryotic genomic DNA is packaged into chromatin through association with positively charged histones to form the nucleosome, the structural unit of chromatin (1–3). The nucleosome core consists of an octamer of histones with two copies of H2A, H2B, H3, and H4, around which ∼146 bp of DNA winds in ∼1.65 left-handed superhelical turns (4). At this level of the DNA packaging, chromatin resembles a beads-on-a-string structure, with the nucleosome core as the beads and the linker DNA between them as the strings (5). At the next level of DNA packaging, H1 histones bind to the linker DNA and the nucleosome to further condense the chromatin structure (6, 7). H1-mediated chromatin condensation plays important roles in cellular functions such as mitotic chromosome architecture and segregation (8), muscle differentiation (9), and regulation of gene expression (10, 11).Linker H1 histones typically are ∼200 amino acid residues in length, with a short N-terminal region, followed by a ∼70–80-amino acid structured globular domain (gH1) and a ∼100-amino acid unstructured C-terminal domain that is highly enriched in Lys residues. H1 stabilizes the nucleosome and facilitates folding of nucleosome arrays into higher-order structures (12–15). gH1 alone confers the same protection from micrococcal nuclease digestion to the nucleosome as the full-length H1 does (16). The N-terminal region of H1 is not important for nucleosome binding (16, 17), whereas the C terminus is required for H1 binding to chromatin in vivo (18, 19) and for the formation of a stem structure of linker DNA in vitro (17, 20, 21).The globular domain structures of avian H5 (22) and budding yeast Hho1 (23), which are both H1 homologs, have been determined at atomic resolution and show similar structures. In addition, numerous studies have indicated that gH1/gH5 binds around the dyad region of the nucleosome (14, 24), leading to many conflicting structural models for how the globular domain of H1/H5 binds to the nucleosome (SI Appendix, Fig. S1) (24–26). These models are divided into two major classes, symmetric and asymmetric, on the basis of the location of gH1/gH5 in the nucleosome. In the symmetric class, gH1/gH5 binds to the nucleosomal DNA at the dyad and interacts with both linker DNAs (16, 17, 27, 28). In the asymmetric class, gH1/gH5 binds to the nucleosomal DNA in the vicinity of the dyad axis and to 10 bp (27, 29–32) or 20 bp (19, 29, 33, 34) of one linker DNA, or is located inside the DNA gyres, where it interacts with histone H2A (35). In addition, Zhou and colleagues also characterized the orientation of gH5 in the gH5-nucleosome complex (29). The use of nonuniquely positioned nucleosomes and indirect methods may have contributed to the differences in these models (SI Appendix, Fig. S1).Multidimensional nuclear magnetic resonance (NMR), and in particular methyl-based NMR, provides a direct approach to the structural characterization of macromolecular complexes (36, 37). We have previously assigned chemical shifts of the methyl groups of the side chains of residues Ile, Leu, and Val in the core histones (38) and the backbone amides in the disordered histone tails (39), which provide the fingerprints for investigating the interactions between H1 and the nucleosome. Here, we used NMR, along with several other methods, to determine the location and orientation of the globular domain of a stable mutant of Drosophila H1 on a well-positioned nucleosome.