George Sirinakis

Single Reconstituted Neuronal SNARE Complexes Zipper in Three Distinct Stages

Dissecting SNARE Zippering The SNARE complex is critical for vesicle fusion, notably during release of neurotransmitters at synapses. Understanding the biophysics of SNARE assembly has been the object of several structural studies, and yet much remains to be understood about the mechanisms. Now, Gao et al. (p. 1340, published online 16 August; see the Perspective by Rizo) describe the results of cell-free experiments using optical tweezers to elucidate assembly and disassembly of the SNARE complex. Direct observations of SNARE intermediates revealed multiple steps of the assembly process, along with the associated energetics and kinetics. Applying forces similar to those occurring during fusion, an intermediate was stabilized, and the derived mechanism indicates how neurotransmitter release may be regulated. Zippering of a single SNARE complex generates high force and energy that can potentially drive synaptic membrane fusion. Soluble N-ethylmaleimide–sensitive factor attachment protein receptor (SNARE) proteins drive membrane fusion by assembling into a four-helix bundle in a zippering process. Here, we used optical tweezers to observe in a cell-free reconstitution experiment in real time a long-sought SNARE assembly intermediate in which only the membrane-distal amino-terminal half of the bundle is assembled. Our findings support the zippering hypothesis, but suggest that zippering proceeds through three sequential binary switches, not continuously, in the amino- and carboxyl-terminal halves of the bundle and the linker domain. The half-zippered intermediate was stabilized by externally applied force that mimicked the repulsion between apposed membranes being forced to fuse. This intermediate then rapidly and forcefully zippered, delivering free energy of 36 kBT (where kB is Boltzmann’s constant and T is temperature) to mediate fusion.

Ultra-High Resolution 3D Imaging of Whole Cells

Summary Fluorescence nanoscopy, or super-resolution microscopy, has become an important tool in cell biological research. However, because of its usually inferior resolution in the depth direction (50–80 nm) and rapidly deteriorating resolution in thick samples, its practical biological application has been effectively limited to two dimensions and thin samples. Here, we present the development of whole-cell 4Pi single-molecule switching nanoscopy (W-4PiSMSN), an optical nanoscope that allows imaging of three-dimensional (3D) structures at 10- to 20-nm resolution throughout entire mammalian cells. We demonstrate the wide applicability of W-4PiSMSN across diverse research fields by imaging complex molecular architectures ranging from bacteriophages to nuclear pores, cilia, and synaptonemal complexes in large 3D cellular volumes.

The RSC chromatin remodelling ATPase translocates DNA with high force and small step size

ATP‐dependent chromatin remodelling complexes use the energy of ATP hydrolysis to reposition and reconfigure nucleosomes. Despite their diverse functions, all remodellers share highly conserved ATPase domains, many shown to translocate DNA. Understanding remodelling requires biophysical knowledge of the DNA translocation process: how the ATPase moves DNA and generates force, and how translocation and force generation are coupled on nucleosomes. Here, we characterize the real‐time activity of a minimal RSC translocase ‘motor’ on bare DNA, using high‐resolution optical tweezers and a ‘tethered’ translocase system. We observe on dsDNA a processivity of ∼35 bp, a speed of ∼25 bp/s, and a step size of 2.0 (±0.4, s.e.m.) bp. Surprisingly, the motor is capable of moving against high force, up to 30 pN, making it one of the most force‐resistant motors known. We also provide evidence for DNA ‘buckling’ at initiation. These observations reveal the ATPase as a powerful DNA translocating motor capable of disrupting DNA–histone interactions by mechanical force.

Two-colour live-cell nanoscale imaging of intracellular targets

Stimulated emission depletion (STED) nanoscopy allows observations of subcellular dynamics at the nanoscale. Applications have, however, been severely limited by the lack of a versatile STED-compatible two-colour labelling strategy for intracellular targets in living cells. Here we demonstrate a universal labelling method based on the organic, membrane-permeable dyes SiR and ATTO590 as Halo and SNAP substrates. SiR and ATTO590 constitute the first suitable dye pair for two-colour STED imaging in living cells below 50 nm resolution. We show applications with mitochondria, endoplasmic reticulum, plasma membrane and Golgi-localized proteins, and demonstrate continuous acquisition for up to 3 min at 2-s time resolution.

Highly Anisotropic Stability and Folding Kinetics of a Single Coiled Coil Protein under Mechanical Tension

Coiled coils are one of the most abundant protein structural motifs and widely mediate protein interactions and force transduction or sensation. They are thus model systems for protein engineering and folding studies, particularly the GCN4 coiled coil. Major single-molecule methods have also been applied to this protein and revealed its folding kinetics at various spatiotemporal scales. Nevertheless, the folding energy and the kinetics of a single GCN4 coiled coil domain have not been well determined at a single-molecule level. Here we used high-resolution optical tweezers to characterize the folding and unfolding reactions of a single GCN4 coiled coil domain and their dependence on the pulling direction. In one axial and two transverse pulling directions, we observed reversible, two-state transitions of the coiled coil in real time. The transitions equilibrate at pulling forces ranging from 6 to 12 pN, showing different stabilities of the coiled coil in regard to pulling direction. Furthermore, the transition rates vary with both the magnitude and the direction of the pulling force by greater than 1000 folds, indicating a highly anisotropic and topology-dependent energy landscape for protein transitions under mechanical tension. We developed a new analytical theory to extract energy and kinetics of the protein transition at zero force. The derived folding energy does not depend on the pulling direction and is consistent with the measurement in bulk, which further confirms the applicability of the single-molecule manipulation approach for energy measurement. The highly anisotropic thermodynamics of proteins under tension should play important roles in their biological functions.

Single-molecule observation of helix staggering, sliding, and coiled coil misfolding

The biological functions of coiled coils generally depend on efficient folding and perfect pairing of their α-helices. Dynamic changes in the helical registry that lead to staggered helices have only been proposed for a few special systems and not found in generic coiled coils. Here, we report our observations of multiple staggered helical structures of two canonical coiled coils. The partially folded structures are formed predominantly by coiled coil misfolding and occasionally by helix sliding. Using high-resolution optical tweezers, we characterized their energies and transition kinetics at a single-molecule level. The staggered states occur less than 2% of the time and about 0.1% of the time at zero force. We conclude that dynamic changes in helical registry may be a general property of coiled coils. Our findings should have broad and unique implications in functions and dysfunctions of proteins containing coiled coils.

Combined versatile high-resolution optical tweezers and single-molecule fluorescence microscopy

Optical trapping and single-molecule fluorescence are two major single-molecule approaches. Their combination has begun to show greater capability to study more complex systems than either method alone, but met many fundamental and technical challenges. We built an instrument that combines base-pair resolution dual-trap optical tweezers with single-molecule fluorescence microscopy. The instrument has complementary design and functionalities compared with similar microscopes previously described. The optical tweezers can be operated in constant force mode for easy data interpretation or in variable force mode for maximum spatiotemporal resolution. The single-molecule fluorescence detection can be implemented in either wide-field or confocal imaging configuration. To demonstrate the capabilities of the new instrument, we imaged a single stretched λ DNA molecule and investigated the dynamics of a DNA hairpin molecule in the presence of fluorophore-labeled complementary oligonucleotide. We simultaneously observed changes in the fluorescence signal and pauses in fast extension hopping of the hairpin due to association and dissociation of individual oligonucleotides. The combined versatile microscopy allows for greater flexibility to study molecular machines or assemblies at a single-molecule level.

DNA Translocation of ATP-Dependent Chromatin Remodeling Factors Revealed by High-Resolution Optical Tweezers

ATP-dependent chromatin remodeling complexes (remodelers) use the energy of ATP hydrolysis to regulate chromatin structures by repositioning and reconfiguring nucleosomes. Ensemble experiments have suggested that remodeler ATPases are DNA translocases, molecular motors capable of processively moving along DNA. This concept of DNA translocation has become a foundation for understanding the molecular mechanisms of ATP-dependent chromatin remodeling and its biological functions. However, quantitative characterizations of DNA translocation by representative remodelers are rare. Furthermore, it is unclear how a unified theory of chromatin remodeling is built upon this foundation. To address these problems, high-resolution optical tweezers have been applied to investigate remodeler translocation on bare DNA and nucleosomal DNA substrates at a single-molecule level. Our strategy is to hold two ends of a single DNA molecule and measure remodeler translocation by detecting the end-to-end extension and tension changes of the DNA molecule in response to chromatin remodeling. These single-molecule assays can reveal detailed kinetics of remodeler translocation, including velocity, processivity, stall force, pauses, direction changes, and even step size. Here we describe instruments, reagents, sample preparations, and detailed protocols for the single-molecule experiments. We show that optical tweezer force microscopy is a powerful and friendly tool for studies of chromatin structures and remodeling.

The RSC Chromatin Remodeling ATPase can Translocate DNA with High Force and Small Step Size

ATP-dependent chromatin remodeling complexes (remodelers) use the energy of ATP hydrolysis to reposition and reconfigure nucleosomes. Despite their diverse functions, all remodelers share highly conserved catalytic ATPase domains, many of which are shown to translocate DNA. Understanding remodeling requires biophysical knowledge of the DNA translocation process: how the ATPase moves DNA and generates force, and how translocation and force generation are coupled on nucleosomes. Here we characterize the real-time activity of a minimal translocase ‘motor’ complex isolated from a prototypical remodeler (RSC) on bare DNA, using high-resolution optical tweezers and a ‘tethered’ translocase system. We observe on dsDNA a processivity of ∼35 bp, a speed of ∼25 bp/sec, and a step size of 1.9 (± 0.3, s.d.) bp. Surprisingly, the motor is capable of moving against high force, up to 30 pN, making it one of the most force-resistant motors known. We also provide evidence for DNA ‘buckling’ at initiation. These observations extend and clarify measurements of nucleosome-dependent translocation by the complete RSC or SWI/SNF complex, and reveal the ATPase as a powerful and versatile DNA translocating motor capable of disrupting DNA-histone interactions by mechanical force using a small step size.