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Chuck Sindelar, PhD

Research Scientist

Contact Information

Chuck Sindelar, PhD

Mailing Address

  • 333 Cedar Street, CE-25 SHM

    P.O. Box 208024

    New Haven, CT 06520-8024

    United States

Research Summary

My group's recent work has turned towards advancing cryo-electron microscopy techniques to the point where atomic-level features can be obtained for systems such as cytoskeletal filaments. These efforts led to our recent synthesis, using cryo-electron microscopy (cryo-EM) and X-ray crystallography information, of an atomic-level model for kinesin’s ATP-sensing machinery in its active form, which is only assumed following microtubule attachment. This discovery led to a simple and intuitive “clamshell” mechanism describing how ATP binding leads to force generation in the microtubule-attached motor. My laboratory's research interests have expanded to include numerous other filament-related molecular machines, such the myosin molecular motor that powers muscle movement and the fascinating endoflagellum that spirochete bacteria to convert their bodies into pathogenic 'drilling machines'.

Extensive Research Description

The mechanism of kinesin motility and force production

I have a longstanding interest in the structural basis of motility by the kinesin cytoskeletal motor proteins. As a dimeric molecule of kinesin ‘walks’ along the microtubule powered by ATP hydrolysis, each kinesin motor domain must periodically attach and detach from the filament surface, closely orchestrated with its partner domain– a fantastically intricate process that remains inadequately explained by existing structural models. This pursuit led me to cryo-electron microscopy– the only structural method able to visualize these motors while attached to their microtubule tracks. Beginning with my postdoctoral studies with Ken Downing and Niko Grigorieff and continuing after I started my own research laboratory, I used cryo-EM to produce the first accurate atomic models capturing the sequence of structural changes undergone by the monomeric kinesin motor domain as it binds to the microtubule, releases ADP, then binds and hydrolyses ATP (Sindelar and Downing, JCB, 2007); Sindelar and Downing, PNAS, 2010; Shang et al., eLife, 2013). These studies revealed that microtubules remodel kinesin’s structure to enable a previously unobserved ‘clamshell’ behavior, powered by a twisting of the entire molecule similar to a wind-up toy. A subsequent technical breakthrough by our group enabled us to directly visualize, for the first time, fully active dimeric kinesin poised in mid-step along the microtubule (Liu et al., eLife, 2017), permitting direct testing of structural motility models arising from our prior work with monomeric kinesins.

Building on the above structural studies, we have now developed a new ‘patch’ method (Debs et al., PNAS 2020) that greatly improves on our earlier cryo-EM methods by adapting state-of-the-art maximum-likelihood refinement and structural classification techniques to specifically account for asymmetric features of helical assemblies. With this method, we obtained a structure of the microtubule stabilized by a cancer drug, taxol, at 2.9Å resolution(ibid) – a considerable advance over the best resolution previously achieved with similar sample. The method has also yielded a new structure of the ‘empty’ state of monomeric kinesin-1 bound to an intact microtubule, which provides the first direct validation for the structure models of this state suggested by our earlier cryo-EM studies at lower resolution. Moreover, the patch method enabled us to capture a series of key structural intermediates describing a forward step by the dimeric form of kinesin (manuscript in preparation). By extending these results to near-atomic resolution, we will pursue the ‘holy grail’ of kinesin mechanism– how the energy liberated by ATP hydrolysis and phosphate release couples to the forward step.

Studies relating to the actin cytoskeleton, including the myosin molecular motor

My group at Yale has leveraged the high-resolution structure-determination methods we developed for microtubules in order to study processes related to actin filaments, including myosin motility and actin filament severing by the regulatory factor cofilin. In collaboration with the laboratories of Anne Houdusse and Lee Sweeney, we solved a sub-nanometer reconstruction of the unconventional myosin X motor to reveal an unexpectedly extreme lever arm angle at the end of the power stroke, helping explain this motor’s unique ability to step along actin filament bundles (Ropars et al., Nat. Comm., 2015). We also initiated a collaboration with the laboratory of Mike Ostap to study the structural basis of force-sensitivity in myosin-1b, which recently culminated by our elucidation of this motor’s ADP release pathway at near-atomic resolution by cryo-EM (Mentes et al., PNAS, 2018) These structures revealed a novel interface between the myosin1b lever arm and the upper side of its nucleotide cleft that modulates the force sensitivity. Moreover, by applying novel structural classification methods, we discovered two distinct populations of myo1b with ADP bound, allowing us to present detailed mechanism of force-inhibited ADP release for myo1b that also explains why many myosins lack this type of force sensitivity.

In a separate line of research, we have investigated cytoskeletal factors that control the morphology of actin filaments. We teamed up with the laboratory of Enrique De La Cruz to solve structures of cofilin-bound actin filaments, demonstrating a role for cation binding in the severing mechanism (Kang et al., PNAS, 2013), and also solved the structure of a phosphomimetic mutant that contributed to a model for regulation of cofilin severing (Elam et al., JBC, 2017). In our most recent publications on this topic, we combined state of the art cryo-EM methods with specialized in-house tools for structural classification and filament geometry analysis, to discover an abrupt structural transition, or ‘phase boundary’, in actin filaments initiated by the binding of the cofilin (Huehn et al., JBC, 2018; Huehn et al., PNAS 2020). Our analysis demonstrated that these boundaries, where severing takes place, exhibit abrupt changes in filament geometry, settling an important controversy in the field and paving the way for ongoing work by our group to resolve the boundary structure at high-resolution in 3D. Finally, in collaboration with David Calderwoord we solved the first near-atomic resolution structure of the ABD family of actin cross-linking factors, bound to the actin filament (Iwamoto et al., NSMB, 2018). This work gives a detailed description of the filamin-actin interface and yields the structural basis for a variety of disease-causing mutations.

Molecular architecture of the flagellar filament in spirochete bacteria

The flagellum acts as a rotary propeller that drives bacterial motility and pathogenesis. It is, however, an extremely challenging target for structure determination due to its flexibility, small diameter, and capacity to switch between a multiplicity of different structural states. Only a handful of bacterial flagella have ever been visualized at high resolution, all being mutated forms where motility is compromised. By developing new electron cryo-tomography analysis methods, my group has now solved a novel flagellar structure from a spirochete, a class of bacteria with an internalized flagellum that drives pathogenesis in a variety of spirochete-borne illnesses including syphilis and Lyme disease (Gibson et al., eLife, 2020).

At 10Å resolution, our reconstruction of the Leptospiraflagellum allowed us to build an atomic model for most of the assembly, including two pathogenicity factors that were recently discovered and crystallized by our collaborators (Albert Ko group, Yale Public Health; Alejandro Buschiazzo, Institute Pasteur, Uruguay). Our model reveals a conserved ‘core’ domain surrounded by a radially asymmetric ‘sheath’ domain. This novel arrangement appears to be unique to spirochetes. Over the next five years, a major research goal will be to improve the resolution and extend these studies to the in situcase, aided by further methodological advancements.

SNARE-mediated synaptic vesicle fusion

My group has collaborated closely with laboratory of James Rothman to perform structural studies to elucidate the process of SNARE-mediated vesicle fusion, the process by which vesicles merge with a target membrane to enable intra- and inter-cellular transfer of neurotransmitters and other vital biomolecules (Wang et al., PNAS, 2014; Zanetti et al., eLife, 2016; Wang et al., eLife, 2017; Grushin et al., N. Comm, 2019). This process is tightly coupled to calcium and must occur at very high speed (millisecond timescale) to support effective neurotransmission. Together, our groups discovered that synaptotagmin, an accessory protein vital for functional calcium-triggering of membrane fusion in many cells (including neurons), can oligomerize on membranes into rings and tubular helical assemblies in a calcium-dependent manner. We elaborated on this discovery in a series of publications that have led to the hypothesis that rings of synaptotagmin form the core of a ‘buttressed ring’ that would clamp vesicles in a pre-fusion state, poised for a subsequent, synchronous fusion event triggered by calcium.

Advancing the general theory of cryo-EM image-processing and 3D reconstruction.

Beginning in a second postdoctoral position at Brandeis University and continuing on during my Professorship at Yale University, I became interested in the question of how to optimize the process of merging very noisy images obtained by cryo-EM into a three-dimensional reconstruction. The result was a pair of papers that give an explanation for the perplexing observation that ‘optimizing’ the signal to noise ratio of 2D or 3D image averages using the so-called Wiener filter gives unexpectedly blurry results, when applied to single particle images typically collected in cryo-EM studies (Sindelar and Grigorieff, J. Struct. Biol., 2011; Sindelar and Grigorieff, J. Struct. Biol., 2012). I found that a correction factor had to be introduced into the Wiener filter equation, and derived an analytical formula that relates this correction factor to the size of the particle being imaged. Moreover, I showed that the resulting theoretical framework allows the particle size to be estimated solely from statistical comparisons of noisy images in the cryo-EM data set. The resulting modifications to the Wiener filter have been incorporated into the FREALIGN software package (Sindelar and Grigorieff, J. Struct. Biol. 2012) and forms an integral component of its successor called ‘cisTEM’ as well as a recent sub-tomographic averaging suite (‘emClarity’), two cutting-edge methods for high-resolution 3D refinement and reconstruction of cryo-EM data.


Research Interests

Adenosine Triphosphate; Biochemistry; Biophysics; Kinesin; Crystallography, X-Ray; Cryoelectron Microscopy

Selected Publications