An atomic-level mechanism for activation of the kinesin molecular motors

Kinesin cytoskeletal motors convert the energy of ATP hydrolysis into stepping movement along microtubules. A partial model of this process has been derived from crystal structures, which show that movement of the motor domain relative to its major microtubule binding element, the switch II helix, is coupled to docking of kinesin’s neck linker element along the motor domain. This docking would displace the cargo in the direction of travel and so contribute to a step. However, the crystal structures do not reveal how ATP binding and hydrolysis govern this series of events. We used cryoelectron microscopy to derive 8–9 Å-resolution maps of four nucleotide states encompassing the microtubule-attached kinetic cycle of a kinesin motor. The exceptionally high quality of these maps allowed us to build in crystallographically determined conformations of kinesin’s key subcomponents, yielding novel arrangements of kinesin’s switch II helix and nucleotide-sensing switch loops. The resulting atomic models reveal a seesaw mechanism in which the switch loops, triggered by ATP binding, propel their side of the motor domain down and thereby elicit docking of the neck linker on the opposite side of the seesaw. Microtubules engage the seesaw mechanism by stabilizing the formation of extra turns at the N terminus of the switch II helix, which then serve as an anchor for the switch loops as they modulate the seesaw angle. These observations explain how microtubules activate kinesin’s ATP-sensing machinery to promote cargo displacement and inform the mechanism of kinesin’s ancestral relative, myosin.     

Three-dimensional deconvolution processing for STEM cryotomography

The complex environment of biological cells and tissues has motivated development of three-dimensional (3D) imaging in both light and electron microscopies. To this end, one of the primary tools in fluorescence microscopy is that of computational deconvolution. Wide-field fluorescence images are often corrupted by haze due to out-of-focus light, i.e., to cross-talk between different object planes as represented in the 3D image. Using prior understanding of the image formation mechanism, it is possible to suppress the cross-talk and reassign the unfocused light to its proper source post facto. Electron tomography based on tilted projections also exhibits a cross-talk between distant planes due to the discrete angular sampling and limited tilt range. By use of a suitably synthesized 3D point spread function, we show here that deconvolution leads to similar improvements in volume data reconstructed from cryoscanning transmission electron tomography (CSTET), namely a dramatic in-plane noise reduction and improved representation of features in the axial dimension. Contrast enhancement is demonstrated first with colloidal gold particles and then in representative cryotomograms of intact cells. Deconvolution of CSTET data collected from the periphery of an intact nucleus revealed partially condensed, extended structures in interphase chromatin.

Electron tomography (ET) is the premier technique for visualization of cellular ultrastructure in three dimensions (3D). ET actually encompasses a number of different experimental approaches (1–3). These include serial sections imaged by transmission electron microscopy (TEM) and “serial surface” views produced by scanning electron microscopy (SEM) combined with serial or iterative microtomy. A 3D image is produced by aligning and combining the sections. Most commonly, ET is performed by rotating a specimen around an axis while a series of projection images is recorded by wide-field TEM. A 3D image can then be reconstructed from the projections by standard algorithms (4, 5). ET is routinely performed on plastic embedded cell or tissue specimens that have been stained with heavy metals for contrast enhancement. ET on unstained hydrated specimens at cryogenic temperatures can provide even molecular detail (6, 7).Cryogenic vitrification offers potentially the most faithful preservation of biological cells and other aqueous-based materials (8). Imaging without the benefit of contrast-enhancing stains is highly challenging, however. The combination of weak electron scattering and strict exposure limits to avoid specimen damage results in noisy images. In addition, certain systematic artifacts are inherent to reconstructions from projection tilt series. While in specific cases a cylindrical geometry provides for full rotation around the tilt axis (9, 10), the more typical flat slab-geometry specimens can be tilted only within a certain restricted range. This artifact is known as the “missing wedge”; the missing projections impair the reconstruction. Acquisition of projection images at a limited number of discrete angles also results in missing information for reconstruction (11, 12). Both limitations lead to “ghost” contrast that emanates from sharp features and projects into neighboring planes.Ghost artifacts and the smearing of axial contrast in electron tomography recall out-of-focus blur in fluorescent light microscopy. Fluorescence deconvolution algorithms have reached a degree of maturity, yet advances are also underway (13). In particular, entropy regularization makes a significant improvement in noise suppression (14). Assuming that the 3D image of a point source, i.e., the point spread function (PSF), is known, one might similarly reassign the reconstruction artifacts of electron tomography back to the plane of origin by 3D image deconvolution. In this work we ask whether algorithms intended for optical deconvolution may serve to enhance 3D reconstruction by electron cryotomography.At first sight, the very different configurations of fluorescence and electron microscopies make such an application unlikely. Fluorescence emission behaves as a source of light; its intensity is (within limits) quantitatively proportional to the local fluorophore density in the specimen. Transmission electron microscopy detects electron scattering by the projected electrostatic potential, which is (very) approximately proportional to electron density; the scattering contrast between water and organic materials is inherently low. Fluorescence imaging is typically performed with objective lenses of high numerical aperture with a cylindrically symmetric PSF covering as much as 70° in collection semiangle. For cryoelectron tomography, projection images are normally recorded one by one using parallel illumination and phase contrast in wide-field TEM. Fluorescence contrast is unipolar—bright on a dark background. Phase images are interferometric by nature; they contain areas both brighter and darker than background due to constructive and destructive interference. Reliance on phase contrast also introduces a number of complications specifically for thick specimens. These originate from inelastic scattering and chromatic aberrations, on one hand, and from violation of the weak phase object approximation on the other (15).Tomography by scanning TEM (STEM) was introduced for plastic embedded section (16) and offers an alternative to wide-field phase contrast for cryotomography (17–20). In STEM, a focused probe is scanned across the specimen while independent detectors record the scattered (dark field) and unscattered (bright field) signals. The convergence angle of the illumination cone, as well as the angular ranges collected by the detectors, are under user control (20–22). Under incoherent imaging conditions, contrast depends on differential scattering cross-sections weighted by the densities of constituent atoms and integrated over the detector apertures. For cryoscanning transmission electron tomography (CSTET), a small convergence angle is often used in order to extend the depth of field (18, 23), while dynamic focusing adjusts the focus as a function of specimen tilt and distance of the scanned probe from the tilt axis. Scanned images recorded over a range of specimen tilts can be reconstructed conventionally by back projection. Linearity of the image contrast does not depend on the weak phase limit so the projection assumption for tomography is better satisfied by STEM than by TEM for thick specimens (20). In practice, the limiting thickness may exceed 1 μm.Deconvolution is based on the image of an ideal point source, the PSF. In STEM, incoherent electron scattering in the specimen acts as a local source, similarly to fluorescence emission in light microscopy. Per image, the PSF is approximated simply by the illumination profile, i.e., a diffraction-limited beam focused on the specimen. Since the back-projection operation is essentially additive, an effective 3D PSF can be synthesized as the sum of individual PSFs tilted to the appropriate angles. Rotation around a single axis produces a fan-like pattern in the plane perpendicular. This pattern becomes the 3D PSF for deconvolution of the reconstructed volume (Fig. 1). Three-dimensional deconvolution suppresses artifactual projections that produce a speckling in distant reconstructed planes and reduce contrast in depth. The effect was seen most clearly with colloidal gold particles. Applied to cellular data, contrast enhancement by deconvolution enhances the visibility of low-contrast features as well. Most strikingly it allowed for sharp distinctions between high- and low-density chromatin regions inside the cell nucleus.Open in a separate windowFig. 1.STEM configuration and synthesis of the 3D PSF. A STEM image is formed by scanning a focused electron beam (angles not to scale) across the specimen and collecting the scattered electrons on area detectors. These may include an on-axis bright-field (BF) disk and/or a dark-field annulus (ADF), each of which integrates the scattered flux over a certain angular range. For tomography, a series of projection images is recorded as the specimen is tilted. The boxed Inset shows the construction of the 3D point spread function from a sum of rays representing the illumination profile, tilted to the relevant acquisition angles.     

Neutralization of West Nile virus by cross-linking of its surface proteins with Fab fragments of the human monoclonal antibody CR4354

Many flaviviruses are significant human pathogens, with the humoral immune response playing an essential role in restricting infection and disease. CR4354, a human monoclonal antibody isolated from a patient, neutralizes West Nile virus (WNV) infection at a postattachment stage in the viral life-cycle. Here, we determined the structure of WNV complexed with Fab fragments of CR4354 using cryoelectron microscopy. The outer glycoprotein shell of a mature WNV particle is formed by 30 rafts of three homodimers of the viral surface protein E. CR4354 binds to a discontinuous epitope formed by protein segments from two neighboring E molecules, but does not cause any detectable structural disturbance on the viral surface. The epitope occurs at two independent positions within an icosahedral asymmetric unit, resulting in 120 binding sites on the viral surface. The cross-linking of the six E monomers within one raft by four CR4354 Fab fragments suggests that the antibody neutralizes WNV by blocking the pH-induced rearrangement of the E protein required for virus fusion with the endosomal membrane.     

Monolayer purification: a rapid method for isolating protein complexes for single-particle electron microscopy

Visualizing macromolecular complexes by single-particle electron microscopy (EM) entails stringent biochemical purification, specimen preparation, low-dose imaging, and 3D image reconstruction. Here, we introduce the "monolayer purification" method, which employs nickel-nitrilotriacetic acid (Ni-NTA) functionalized lipids for simultaneously purifying His-tagged complexes directly from cell lysates while producing specimens suitable for single-particle EM. The method was established by using monolayers containing Ni-NTA lipid to specifically adsorb His-tagged transferrin-transferrin receptor (Tf-TfR) complexes from insect and mammalian cell extracts. The specificity and sensitivity of the method could be improved by adding imidazole to the extracts. The monolayer-purified Tf-TfR samples could be vitrified and used to calculate a 3D reconstruction of the complex. Monolayer purification was then used to rapidly isolate ribosomal complexes from bacteria by overexpressing a single His-tagged ribosomal subunit. The resulting monolayer samples allowed calculation of a cryo-EM 3D reconstruction of the Escherichia coli 50S ribosomal subunit.     

Structure of TRPV1 channel revealed by electron cryomicroscopy

The transient receptor potential (TRP) family of ion channels participate in many signaling pathways. TRPV1 functions as a molecular integrator of noxious stimuli, including heat, low pH, and chemical ligands. Here, we report the 3D structure of full-length rat TRPV1 channel expressed in the yeast Saccharomyces cerevisiae and purified by immunoaffinity chromatography. We demonstrate that the recombinant purified TRPV1 channel retains its structural and functional integrity and is suitable for structural analysis. The 19-A structure of TRPV1 determined by using single-particle electron cryomicroscopy exhibits fourfold symmetry and comprises two distinct regions: a large open basket-like domain, likely corresponding to the cytoplasmic N- and C-terminal portions, and a more compact domain, corresponding to the transmembrane portion. The assignment of transmembrane and cytoplasmic regions was supported by fitting crystal structures of the structurally homologous Kv1.2 channel and isolated TRPV1 ankyrin repeats into the TRPV1 structure.     

Surface architecture of endospores of the Bacillus cereus/anthracis/thuringiensis family at the subnanometer scale

Bacteria of the Bacillus cereus family form highly resistant spores, which in the case of the pathogen B. anthracis act as the agents of infection. The outermost layer, the exosporium, enveloping spores of the B. cereus family as well as a number of Clostridia, plays roles in spore adhesion, dissemination, targeting, and germination control. We have analyzed two naturally crystalline layers associated with the exosporium, one representing the "basal" layer to which the outermost spore layer ("hairy nap") is attached, and the other likely representing a subsurface ("parasporal") layer. We have used electron cryomicroscopy at a resolution of 0.8-0.6 nm and circular dichroism spectroscopic measurements to reveal a highly α-helical structure for both layers. The helices are assembled into 2D arrays of "cups" or "crowns." High-resolution atomic force microscopy of the outermost layer showed that the open ends of these cups face the external environment and the highly immunogenic collagen-like fibrils of the hairy nap (BclA) are attached to this surface. Based on our findings, we present a molecular model for the spore surface and propose how this surface can act as a semipermeable barrier and a matrix for binding of molecules involved in defense, germination control, and other interactions of the spore with the environment.     

Skeletal muscle resists stretch by rapid binding of the second motor domain of myosin to actin

A shortening muscle is a machine that converts metabolic energy into mechanical work, but, when a muscle is stretched, it acts as a brake, generating a high resistive force at low metabolic cost. The braking action of muscle can be activated with remarkable speed, as when the leg extensor muscles rapidly decelerate the body at the end of a jump. Here we used time-resolved x-ray and mechanical measurements on isolated muscle cells to elucidate the molecular basis of muscle braking and its rapid control. We show that a stretch of only 5 nm between each overlapping set of myosin and actin filaments in a muscle sarcomere is sufficient to double the number of myosin motors attached to actin within a few milliseconds. Each myosin molecule has two motor domains, only one of which is attached to actin during shortening or activation at constant length. A stretch strains the attached motor domain, and we propose that combined steric and mechanical coupling between the two domains promotes attachment of the second motor domain. This mechanism allows skeletal muscle to resist external stretch without increasing the force per motor and provides an answer to the longstanding question of the functional role of the dimeric structure of muscle myosin.     

The structure of the native cardiac thin filament at systolic Ca2+ levels

Every heartbeat relies on cyclical interactions between myosin thick and actin thin filaments orchestrated by rising and falling Ca²⁺ levels. Thin filaments are comprised of two actin strands, each harboring equally separated troponin complexes, which bind Ca²⁺ to move tropomyosin cables away from the myosin binding sites and, thus, activate systolic contraction. Recently, structures of thin filaments obtained at low (pCa ∼9) or high (pCa ∼3) Ca²⁺ levels revealed the transition between the Ca²⁺-free and Ca²⁺-bound states. However, in working cardiac muscle, Ca²⁺ levels fluctuate at intermediate values between pCa ∼6 and pCa ∼7. The structure of the thin filament at physiological Ca²⁺ levels is unknown. We used cryoelectron microscopy and statistical analysis to reveal the structure of the cardiac thin filament at systolic pCa = 5.8. We show that the two strands of the thin filament consist of a mixture of regulatory units, which are composed of Ca²⁺-free, Ca²⁺-bound, or mixed (e.g., Ca²⁺ free on one side and Ca²⁺ bound on the other side) troponin complexes. We traced troponin complex conformations along and across individual thin filaments to directly determine the structural composition of the cardiac native thin filament at systolic Ca²⁺ levels. We demonstrate that the two thin filament strands are activated stochastically with short-range cooperativity evident only on one of the two strands. Our findings suggest a mechanism by which cardiac muscle is regulated by narrow range Ca²⁺ fluctuations.

Striated muscle contraction and relaxation is governed by rising and falling Ca²⁺ levels that control actomyosin interactions—the force generating interaction between the thick (myosin) and thin (actin) filaments (1–4). Thin filaments (TFs) are comprised of F-actin, the troponin complex (Tn), and tropomyosin (Tm). Tn itself is a multiprotein complex comprised of the TnI (e.g., inhibitory) subunit, the TnC (e.g., Ca²⁺ binding) subunit, and TnT—the subunit which links the Tn complex to the Tm cable. During muscle activation, conformational changes within Tn caused by Ca²⁺ binding to its TnC subunit allosterically relieve the Tm-induced actomyosin inhibition (4–6). Kinetics, biochemical, and structural studies led to the steric blocking model of muscle regulation, which holds that at low Ca²⁺ (relaxing) conditions, Tm blocks myosin binding sites on the actin surface, whereas at high Ca²⁺ (activating) conditions, Tm moves away from the myosin binding sites (5, 7). Early electron microscopy (EM) studies of negatively stained TFs identified three structural states for Tm, termed “blocked,” “closed,” and “myosin” (“open”), which corresponded to relaxed, activated, and myosin-bound conditions, respectively (8). Later, structural analysis of frozen hydrated cardiac TFs (9) updated the positioning of the Tm on the surface of the TF and showed that at either relaxing or activating conditions, Tm was not constrained in one structural state but rather was distributed among the three structural positions on the surface of the TF. In all aforementioned studies, a helical approach to three-dimensional (3D) reconstruction was applied to the TFs since the actin filament, which comprises the backbone of the TF, is a helical structure (10). On the other hand, the periodicity of the TF regulatory units (RUs) comprised of one Tm and one Tn complex bound to seven actin protomers along the actin filament is not consistent with the symmetry of the actin helix. For that reason, the helical approach to 3D reconstruction of the TF generates helically averaged Tm densities while eliminating any information on the structure of the Tn complex. Despite those shortcomings, the helical approach to frozen hydrated TFs accurately predicted the range of Tm swing upon activation of the TF and showed a rocking movement of the Tn on the surface of the TF (9). The revolutionary work by Yamada et al. (6) utilized a nonhelical algorithm for reconstruction of the cardiac TF and for the first time ever showed structural transition of the TF RUs from Ca²⁺ free to the Ca²⁺-bound state. While this important study enormously advanced our understanding of the TF regulation, the use of recombinant proteins expressed in a heterologous system without the naturally occurring posttranslational modifications found in the mammalian heart left some questions regarding how accurately the structures represented native cardiac TFs. Furthermore, the structures were obtained in a nonphysiological either Ca²⁺-free (pCa ∼9) (subdiastolic) or excessive (pCa ∼3) suprasystolic Ca²⁺ levels. However, in the working heart, the intracellular Ca²⁺ concentration fluctuates within a range of 0.1 (pCa ∼7) to 1.0 µM (pCa ∼6) during the cardiac cycle (11). Therefore, to understand the mechanism(s) of the TF activation, it was imperative to perform structural studies at physiological Ca²⁺ levels.Here, we used native cardiac TFs to carry out cryogenic EM (cryo-EM) structural studies at systolic (pCa = 5.8) Ca²⁺ levels. The cryo-EM structures and the use of statistical tools in analysis of individual frozen hydrated native cardiac TFs allowed us to develop a model for the structural state of the cardiac TF at systolic Ca²⁺ levels. We demonstrate that the cardiac TF has a network of weakly correlated structural states and operates as a stochastic Ca²⁺-controlled machine.     

Three-dimensional structure of cytoplasmic dynein bound to microtubules

Cytoplasmic dynein is a large, microtubule-dependent molecular motor (1.2 MDa). Although the structure of dynein by itself has been characterized, its conformation in complex with microtubules is still unknown. Here, we used cryoelectron microscopy (cryo-EM) to visualize the interaction between dynein and microtubules. Most dynein molecules in the nucleotide-free state are bound to the microtubule in a defined conformation and orientation. A 3D image reconstruction revealed that dynein's head domain, formed by a ring-like arrangement of AAA+ domains, is located ≈280 Å away from the center of the microtubule. The order of the AAA+ domains in the ring was determined by using recombinant markers. Furthermore, a 3D helical image reconstruction of microtubules with a dynein's microtubule binding domain [dynein stalk (DS)] revealed that the stalk extends perpendicular to the microtubule. By combining the 3D maps of the dynein-microtubule and DS-microtubule complexes, we present a model for how dynein in the nucleotide-free state binds to microtubules and discuss models for dynein's power stroke.     

Architecture of the mycobacterial succinate dehydrogenase with a membrane-embedded Rieske FeS cluster

Complex II, also known as succinate dehydrogenase (SQR) or fumarate reductase (QFR), is an enzyme involved in both the Krebs cycle and oxidative phosphorylation. Mycobacterial Sdh1 has recently been identified as a new class of respiratory complex II (type F) but with an unknown electron transfer mechanism. Here, using cryoelectron microscopy, we have determined the structure of Mycobacterium smegmatis Sdh1 in the presence and absence of the substrate, ubiquinone-1, at 2.53-Å and 2.88-Å resolution, respectively. Sdh1 comprises three subunits, two that are water soluble, SdhA and SdhB, and one that is membrane spanning, SdhC. Within these subunits we identified a quinone-binding site and a rarely observed Rieske-type [2Fe-2S] cluster, the latter being embedded in the transmembrane region. A mutant, where two His ligands of the Rieske-type [2Fe-2S] were changed to alanine, abolished the quinone reduction activity of the Sdh1. Our structures allow the proposal of an electron transfer pathway that connects the substrate-binding and quinone-binding sites. Given the unique features of Sdh1 and its essential role in Mycobacteria, these structures will facilitate antituberculosis drug discovery efforts that specifically target this complex.

During respiration, cells harvest energy from their environment via redox reactions. The harvested energy is converted into adenosine triphosphate (ATP) by ATP synthase (also called as complex V). This process requires the formation of a transmembrane electrochemical gradient that is produced by the electron flux catalyzed by four integral-membrane respiratory complexes (designated complexes I to IV) of the oxidative phosphorylation system (1).Respiratory complex II, also named succinate dehydrogenase (succinate:quinone reductase or SQR) or fumarate reductase (quinol:fumarate reductase or QFR), depending on the preferred direction of the reaction in vivo, performs the reversible oxidation of succinate to fumarate, a function that is coupled to the presence of mobile quinone electron carriers (2). This process is central to cellular metabolism and energy conversion, bridging the Krebs cycle and oxidative phosphorylation (3). All complex II enzymes across the different kingdoms of life share a common overall architecture with a smaller membrane-bound domain and a large soluble domain (4). The soluble domain consists of two hydrophilic subunits, A and B. The structure of subunit A is composed of four domains (5, 6): an FAD-binding domain, a capping domain, a helical domain, and a C-terminal domain. The active site for succinate-fumarate interconversion is located between the flavin-binding domain and the capping domain. Subunit B is a small iron-sulfur protein harboring a [2Fe-2S], a [4Fe-4S], and a [3Fe-4S] cluster, that facilitate electron movement. The two soluble subunits, A and B, are highly conserved across bacteria and mammals. In contrast, the structure and components of the integral-membrane domain, SdhC, can vary containing 0, 1, or 2 membrane-bound subunits with 5 or 6 transmembrane helices and possess varying numbers of heme b groups (0, 1, or 2). Variations also occur according to the type of quinone they use (ubiquinone or menaquinone) and the number of quinone-binding sites (7, 8). As a result, the membrane-spanning regions have distinct evolutionary origins. Thus, the complex II superfamily has been further divided into five subfamilies (types A through E) depending on their biophysical properties (8). Several structures of complex II superfamily enzymes have been determined: type A (exemplified by the Mycobacterium smegmatis (Msm) Sdh2) (9), type B (exemplified by the Wolinella succinogenes QFR) (6), type C [exemplified by the Escherichia coli (10) and porcine (5) SQRs], and type D (exemplified by the E. coli QFR) (11). However, Sdh1 has only recently been identified in mycobacteria and is suggested to represent a new class of respiratory complex II referred to as type F (12). It has been shown that Sdh1 activity is essential for Mycobacterium tuberculosis to adapt to optimal growth and survival under aerobic conditions (13–15). It may accomplish the oxidation of succinate by using higher potential quinones (12, 16). It is suggested that there are enough structural differences between mycobacterial Sdh1 and mitochondrial SQR for anti-TB drug discovery and design (17).In the present study, cryoelectron microscopy (cryo-EM) structures of the M. smegmatis Sdh1 in the presence and absence of substrate, ubiquinone-1, have been determined. These allow the visualization of the electron transport path of a type F complex II system.     

Structural kinetics of myosin by transient time-resolved FRET

For many proteins, especially for molecular motors and other enzymes, the functional mechanisms remain unsolved due to a gap between static structural data and kinetics. We have filled this gap by detecting structure and kinetics simultaneously. This structural kinetics experiment is made possible by a new technique, (TR)(2)FRET (transient time-resolved FRET), which resolves protein structural states on the submillisecond timescale during the transient phase of a biochemical reaction. (TR)(2)FRET is accomplished with a fluorescence instrument that uses a pulsed laser and direct waveform recording to acquire an accurate subnanosecond time-resolved fluorescence decay every 0.1 ms after stopped flow. To apply this method to myosin, we labeled the force-generating region site specifically with two probes, mixed rapidly with ATP to initiate the recovery stroke, and measured the interprobe distance by (TR)(2)FRET with high resolution in both space and time. We found that the relay helix bends during the recovery stroke, most of which occurs before ATP is hydrolyzed, and two structural states (relay helix straight and bent) are resolved in each nucleotide-bound biochemical state. Thus the structural transition of the force-generating region of myosin is only loosely coupled to the ATPase reaction, with conformational selection driving the motor mechanism.     

Translocation pathway of protein substrates in ClpAP protease

Intracellular protein degradation, which must be tightly controlled to protect normal proteins, is carried out by ATP-dependent proteases. These multicomponent enzymes have chaperone-like ATPases that recognize and unfold protein substrates and deliver them to the proteinase components for digestion. In ClpAP, hexameric rings of the ClpA ATPase stack axially on either face of the ClpP proteinase, which consists of two apposed heptameric rings. We have used cryoelectron microscopy to characterize interactions of ClpAP with the model substrate, bacteriophage P1 protein, RepA. In complexes stabilized by ATPgammaS, which bind but do not process substrate, RepA dimers are seen at near-axial sites on the distal surface of ClpA. On ATP addition, RepA is translocated through approximately 150 A into the digestion chamber inside ClpP. Little change is observed in ClpAP, implying that translocation proceeds without major reorganization of the ClpA hexamer. When translocation is observed in complexes containing a ClpP mutant whose digestion chamber is already occupied by unprocessed propeptides, a small increase in density is observed within ClpP, and RepA-associated density is also seen at other axial sites. These sites appear to represent intermediate points on the translocation pathway, at which segments of unfolded RepA subunits transiently accumulate en route to the digestion chamber.     

From the Cover: PNAS Plus: Calcium can mobilize and activate myosin-VI

The ability to coordinate the timing of motor protein activation lies at the center of a wide range of cellular motile processes including endocytosis, cell division, and cancer cell migration. We show that calcium dramatically alters the conformation and activity of the myosin-VI motor implicated in pivotal steps of these processes. We resolved the change in motor conformation and in structural flexibility using single particle analysis of electron microscopic data and identified interacting domains using fluorescence spectroscopy. We discovered that calcium binding to calmodulin increases the binding affinity by a factor of 2,500 for a bipartite binding site on myosin-VI. The ability of calcium-calmodulin to seek out and bridge between binding site components directs a major rearrangement of the motor from a compact dormant state into a cargo binding primed state that is nonmotile. The lack of motility at high calcium is due to calmodulin switching to a higher affinity binding site, which leaves the original IQ-motif exposed, thereby destabilizing the lever arm. The return to low calcium can either restabilize the lever arm, required for translocating the cargo-bound motors toward the center of the cell, or refold the cargo-free motors into an inactive state ready for the next cellular calcium flux.In human cells, cytoskeletal motor proteins move along microtubules and actin filaments to generate complex cellular functions that require a precise timing of motor activation and inactivation. Myosin-VI is thought to have unique properties because it is the only myosin in the human genome shown to move toward the minus end of actin filaments (1). Apart from its roles in the formation of stereocilia in cells of the auditory system (2, 3), membrane internalization (4–6), and delivery of membrane to the leading edge in migratory cells (7), myosin-VI is an early marker of cancer development, aggressiveness, and cancer–cell invasion because of its dramatically up-regulated expression in breast, lung, prostate, ovary, and gastresophagus carcinoma cells (7–11). How this motor might promote cancer–cell migration, proliferation, and survival is unknown.In migrating cells, localized calcium transients (∼50 nM to ∼10 μM) (12, 13) have been reported to play a multifunctional role in steering directional movement (14), cytoskeleton redistribution, and relocation of focal adhesions (15). The effect of calcium transients on the mobilization and cargo binding of myosin-VI and on its mechanical activation, however, are not understood. In the current model, the catalytic head domain hydrolyzes ATP, whereas the tail domain anchors the motor to specific compartments. In vitro studies have shown that calcium affects myosin-VI binding to phospholipids (6), as well as the kinetics and motility rate of the motor (16, 17). The underlying molecular mechanisms, however, are unknown. It has also been discussed that myosin-VI might be able to adopt an inactive folded state (18, 19), perhaps similar to nonmuscle myosin II and myosin-V (20–22), with folding and unfolding regulated by some unknown mechanism. When activated, the myosin-VI head domain binds to actin, generating conformational changes that are transduced by the converter to the lever arm or neck domain and amplified to nanometer displacements. The neck consists of an extended α-helix stabilized by the binding of calmodulin (23), which pointed to the intriguing possibility that the calcium sensor calmodulin bound to the myosin-VI neck domain might constitute a molecular mechanism to control both the cellular mobilization and activation of myosin-VI in migrating cells. We therefore set out to investigate the effect of calcium on the structural conformation, mechanical properties, and activation of single myosin-VI motor molecules using electron microscopy (EM), spectroscopic, and mechanical experiments.     

Direct evidence for functional smooth muscle myosin II in the 10S self-inhibited monomeric conformation in airway smooth muscle cells

The 10S self-inhibited monomeric conformation of myosin II has been characterized extensively in vitro. Based upon its structural and functional characteristics, it has been proposed to be an assembly-competent myosin pool in equilibrium with filaments in cells. It is known that myosin filaments can assemble and disassemble in nonmuscle cells, and in some smooth muscle cells, but whether or not the disassembled pool contains functional 10S myosin has not been determined. Here we address this question using human airway smooth muscle cells (hASMCs). Using two antibodies against different epitopes on smooth muscle myosin II (SMM), two distinct pools of SMM, diffuse, and stress-fiber-associated, were visualized by immunocytochemical staining. The two SMM pools were functional in that they could be interconverted in two ways: (i) by exposure to 10S- versus filament-promoting buffer conditions, and (ii) by exposure to a peptide that shifts the filament-10S equilibrium toward filaments in vitro by a known mechanism that requires the presence of the 10S conformation. The effect of the peptide was not due to a trivial increase in SMM phosphorylation, and its specificity was demonstrated by use of a scrambled peptide, which had no effect. Based upon these data, we conclude that hASMCs contain a significant pool of functional SMM in the 10S conformation that can assemble into filaments upon changing cellular conditions. This study provides unique direct evidence for the presence of a significant pool of functional myosin in the 10S conformation in cells.     

Nonmuscle myosin II exerts tension but does not translocate actin in vertebrate cytokinesis

During vertebrate cytokinesis it is thought that contractile ring constriction is driven by nonmuscle myosin II (NM II) translocation of antiparallel actin filaments. Here we report in situ, in vitro, and in vivo observations that challenge this hypothesis. Graded knockdown of NM II in cultured COS-7 cells reveals that the amount of NM II limits ring constriction. Restoration of the constriction rate with motor-impaired NM II mutants shows that the ability of NM II to translocate actin is not required for cytokinesis. Blebbistatin inhibition of cytokinesis indicates the importance of myosin strongly binding to actin and exerting tension during cytokinesis. This role is substantiated by transient kinetic experiments showing that the load-dependent mechanochemical properties of mutant NM II support efficient tension maintenance despite the inability to translocate actin. Under loaded conditions, mutant NM II exhibits a prolonged actin attachment in which a single mechanoenzymatic cycle spans most of the time of cytokinesis. This prolonged attachment promotes simultaneous binding of NM II heads to actin, thereby increasing tension and resisting expansion of the ring. The detachment of mutant NM II heads from actin is enhanced by assisting loads, which prevent mutant NM II from hampering furrow ingression during cytokinesis. In the 3D context of mouse hearts, mutant NM II-B R709C that cannot translocate actin filaments can rescue multinucleation in NM II-B ablated cardiomyocytes. We propose that the major roles of NM II in vertebrate cell cytokinesis are to bind and cross-link actin filaments and to exert tension on actin during contractile ring constriction.     

Molecular basis of multistep voltage activation in plant two-pore channel 1

Voltage-gated ion channels confer excitability to biological membranes, initiating and propagating electrical signals across large distances on short timescales. Membrane excitation requires channels that respond to changes in electric field and couple the transmembrane voltage to gating of a central pore. To address the mechanism of this process in a voltage-gated ion channel, we determined structures of the plant two-pore channel 1 at different stages along its activation coordinate. These high-resolution structures of activation intermediates, when compared with the resting-state structure, portray a mechanism in which the voltage-sensing domain undergoes dilation and in-membrane plane rotation about the gating charge–bearing helix, followed by charge translocation across the charge transfer seal. These structures, in concert with patch-clamp electrophysiology, show that residues in the pore mouth sense inhibitory Ca²⁺ and are allosterically coupled to the voltage sensor. These conformational changes provide insight into the mechanism of voltage-sensor domain activation in which activation occurs vectorially over a series of elementary steps.

Voltage-gated ion channels (VGICs) use voltage-sensing domains (VSDs) to sense changes in electrical potential across biological membranes (1, 2). VSDs are composed of a four-helix bundle, in which one helix carries charged residues that move in response to changes in transmembrane electric field (3, 4). VSDs usually adopt a “resting state” when the membrane is at “resting potential”: ∼ −80 mV for animal and ∼ −150 mV for plant plasma membranes (5). In comparison, much lower resting membrane voltages are set for intracellular endo-membranes: ∼ −30 mV across the plant vacuole (6) and mammalian lysosome (7). As the membrane potential vanishes during depolarization, so does the downward electrostatic force on the cationic side chains causing them to relax toward the outside of the membrane across a hydrophobic constriction site (HCS) or hydrophobic seal (8). This conformational change is conveyed to the central pore, formed by four pore domains in a quasi-fourfold arrangement, which dilates to allow the diffusion of ions down their electrochemical gradients. The exact nature of the conformational change in VSDs has been the subject of decades of biophysical investigation (9–12), though to this date, only a few structural examples exist of voltage sensors in resting or multiple conformations (13–18).Two-pore channels (TPCs) are defined by their two tandem Shaker-like cassette subunits in a single polypeptide chain, which dimerize to form a C2-symmetric channel with four subunits and 24 (4 × 6) transmembrane helices (19–21). There are three TPC channels, TPC1, 2, and 3, each with different voltage or ligand gating and ion selectivity. Among the voltage-gated TPCs (all except lipid-gated TPC2), only the second VSD (VSD2) is electrically active (18, 22–24), while VSD1 is insensitive to voltage changes and is likely static under all changes in potential.In plants, the vacuole comprises up to 90% of the plant cell volume and provides for a dynamic storage organelle that, in addition to metabolites, is a repository for ions including Ca²⁺. TPC1 channels confer excitability to this intracellular organelle (25) and, unlike other TPCs, are calcium regulated: external Ca²⁺ (in the vacuolar lumen) inhibits the channel by binding to multiple luminal sites, while cytosolic Ca²⁺ is required to open the channel by binding to EF hands, although the exact mechanism by which this activation occurs is unknown (22). These electrical properties allowed our group and Youxing Jiang’s group to determine the first structure of an electrically resting VGIC by cocrystallizing the channel with 1 mM Ca²⁺, which maintains the VSD in a resting configuration at 0 mV potential (16, 22).Previously (17), we used a gain-of-function mutant of AtTPC1 with three luminal Ca²⁺-binding acidic residues on VSD2 neutralized (D240N/D454N/E528N) termed AtTPC1_DDE (abbreviated here as DDE) to visualize channel activation at the level of atomic structure, but we were unable to sufficiently resolve details of the electrically active VSD2 due to structural heterogeneity. In addition, the intracellular activation gate remained closed. We now present multiple structures of intermediately activated states of AtTPC1 determined by extensive image processing. In order to visualize such states, we modulated the channel’s luminal Ca²⁺ sensitivity using a well-studied gain-of-function single-point mutant, D454N (fou2), and also the triple mutant DDE for comparison. fou2 is known to desensitize the channel to inhibitory, external (luminal) calcium ions (26, 27). Mutations in D454 and closely related luminal Ca²⁺-binding carboxyls to alanine D240A, D454A, E528A (termed AtTPC1ΔCa_i) were previously shown to effectively attenuate the Ca²⁺-induced shift of the voltage activation threshold of AtTPC1 to depolarizing potentials at high luminal Ca²⁺ (28).The fou2 and DDE mutations lie in the coordination sphere of the inhibitory Ca²⁺ site on the luminal side of the VSD2–pore interface formed by D454, D240, and E528. The D454N mutation in the fou2 channel enhances the defense capacity of plants against fungal or herbivore attack due to increased production of the wounding hormone jasmonate (29). These effects on plant performance and defense are probably due to short circuiting of the vacuolar membrane (26, 27, 30) in which TPC1 has increased open probability at resting potential. Compared to wild-type (WT) TPC1, the activation threshold in the fou2 channel is shifted to more negatively polarized potentials and also has significantly lower sensitivity to inhibitory Ca²⁺ in addition to exhibiting faster activation kinetics than its WT counterpart that was originally named the “slow vacuolar” (SV) channel due to its slow conductance onset (20, 21, 30). Therefore, D454N confers more than just reduced sensitivity to external Ca²⁺ but intrinsic hyperactivity as well. Our structures of these AtTPC1 mutants attempt to explain how the voltage sensor functions during electrical activation and how exactly luminal Ca²⁺ affects this process.     

Direct structural insight into the substrate-shuttling mechanism of yeast fatty acid synthase by electron cryomicroscopy

Yeast fatty acid synthase (FAS) is a 2.6-MDa barrel-shaped multienzyme complex, which carries out cyclic synthesis of fatty acids. By electron cryomicroscopy of single particles we obtained a three-dimensional map of yeast FAS at 5.9-Å resolution. Compared to the crystal structures of fungal FAS, the EM map reveals major differences and new features that indicate a considerably different arrangement of the complex in solution compared to the crystal structures, as well as a high degree of variance inside the barrel. Distinct density regions in the reaction chambers next to each of the catalytic domains fitted the substrate-binding acyl carrier protein (ACP) domain. In each case, this resulted in the expected distance of ∼18 Å from the ACP substrate-binding site to the active site of the catalytic domains. The multiple, partially occupied positions of the ACP within the reaction chamber provide direct structural insight into the substrate-shuttling mechanism of fatty acid synthesis in this large cellular machine.     

The equilibrium intrinsic crystal–liquid interface of colloids

We use confocal microscopy to study an equilibrated crystal–liquid interface in a colloidal suspension. Capillary waves roughen the surface, but locally the intrinsic interface is sharply defined. We use local measurements of the structure and dynamics to characterize the intrinsic interface, and different measurements find slightly different widths of this interface. In terms of the particle diameter d, this width is either 1.5d (based on structural information) or 2.4d (based on dynamics), both not much larger than the particle size. This work is the first direct experimental visualization of an equilibrated crystal–liquid interface.     

Structural dependence of HET-s amyloid fibril infectivity assessed by cryoelectron microscopy

HET-s is a prion protein of the fungus Podospora anserina which, in the prion state, is active in a self/nonself recognition process called heterokaryon incompatibility. Its prionogenic properties reside in the C-terminal “prion domain.” The HET-s prion domain polymerizes in vitro into amyloid fibrils whose properties depend on the pH of assembly; above pH 3, infectious singlet fibrils are produced, and below pH 3, noninfectious triplet fibrils. To investigate the correlation between structure and infectivity, we performed cryo-EM analyses. Singlet fibrils have a helical pitch of approximately 410 Å and a left-handed twist. Triplet fibrils have three protofibrils whose lateral dimensions (36 × 25 Å) and axial packing (one subunit per 9.4 Å) match those of singlets but differ in their supercoiling. At 8.5-Å resolution, the cross-section of the singlet fibril reconstruction is largely consistent with that of a β-solenoid model previously determined by solid-state NMR. Reconstructions of the triplet fibrils show three protofibrils coiling around a common axis and packed less tightly at pH 3 than at pH 2, eventually peeling off. Taken together with the earlier observation that fragmentation of triplet fibrils by sonication does not increase infectivity, these observations suggest a novel mechanism for self-propagation, whereby daughter fibrils nucleate on the lateral surface of singlet fibrils. In triplets, this surface is occluded, blocking nucleation and thereby explaining their lack of infectivity.