A stepping trace of yeast cytoplasmic dynein labeled with QD-585 and QD-655 shows that the heads step independently of each other, a mechanism fundamentally distinct from coordinated hand-over-hand stepping of kinesin and myosin motors (figure from Dewitt et al., Science 2012).
High-Res Fluorescence Tracking
In conventional light microscopy, the image of a point-like object is a diffraction-limited spot, which has a width of half the wavelength of the emitted light (~250 nm for visible emission). The center of the spot, which represents the position of the probe, can in principle be localized with an arbitrarily high precision by collecting sufficient number of photons from a fluorophore. By collecting more than 10,000 photons per frame, single molecules can be tracked at 1 nm precision. Time resolution of this method is typically 30 ms for organic dyes (Cy3, TMR, Cy5) and a few ms for quantum dot probes. The technique is named Fluorescence Imaging with One Nanometer Accuracy (FIONA).
Using this method, we track the motility of single motors at high precision. These experiments reveal the size and direction of individual steps taken by the motor. By labeling the two heads with a different color, we also determine how the heads are positioned and move relative to each other during processive runs.
FIONA. A. The Airy pattern of a diffraction-limited-spot in two dimensions. B. Fluorescence images of several single Cy3-DNA molecules immobilized on a glass surface. The data was taken with TIRF scope in 0.5 sec. C. Expanded view of one PSF with 2-D elliptical Gaussian curve fit (solid lines).Â The center of this PSF can be located to within 1.5 nm.
Reflection of a well-aligned trapping laser off the bottom surface of the coverslip.
Differential detection stage. Splitting the trapping laser beam into orthogonal polarizations allows us to create two traps simultaneously and detect the positions of two trapped beads independently from one another. Differential measurement experiments allow us to de-couple the observed signal from noise sources common to both trapping beams, such as vibrations of the microscope body or stage drift.
Single-molecule fluorescence resonance energy transfer (smFRET) is a powerful technique that enables us to observe interactions between proteins and nucleic acids, the structure of these complexes as well as their dynamic conformational changes at single molecule level. We label the samples site-specifically with a pair of fluorophores (called donor-acceptor pair) such that the emission spectrum of the donor overlaps with the excitation spectrum of the donor. Because of this overlap, when the donor is brought to its excited state with a laser, the excitation can be transferred to the acceptor, resulting in the emission of an acceptor photon. Since FRET efficiency is a function of the donor-acceptor separation, we can measure distances in the 2-10 nm range on a time scale ranging from milliseconds to several minutes.
Anti-correlated donor and acceptor signals both attached to a DNA molecule and the calculated FRET efficiency. Dynamic conformational changes can be observed.
Non-invasive imaging using fluorescent tagging of various cellular organelles in live or fixed cells is a widely used technique in Cell Biology. However, conventional fluorescence microscopy limits us to a spatial resolution of ~200 nm because of diffraction of light. Hence, many sub-diffraction sized organelles and processes are too small to be imaged with high enough resolution. The details obtained by higher resolution imaging can provide important structural details about these organelles in their functioning. We circumvent the limits of diffraction of light by imaging single molecule in 3 dimensions with improved spatial resolution of ~20 nm using Photoactivable Light Microscopy. In this technique, the proteins that are involved formation of cellular organelles are labeled with a photoactivable GFP ( mEos2, Dronpa etc.). Each labeled protein molecule is then stochastically photoswitched or photoactivated and their location in 3D is determined. The final image is reconstructed from all the molecules that localized giving a super-resolved image.
By tightly focusing a 1064nm laser beam, we can apply piconewton forces to single biological molecules while simultaneously measuring sub-nanometer position and fluctuations at >20 kHz. To appreciate how small the forces applied by molecular motors are, consider that one piconewton is roughly the force of gravitational attraction between two average-sized people standing half a mile apart. Our optical trap is built around a modified Nikon Ti-E microscope and incorporates a custom z-focus feedback system that effectively eliminates axial drift. The trap is built in an acoustically quiet and temperature-controlled room to minimize vibrational noise factors − when measuring molecular-scale motions, someone simply closing a door on the other end of the hallway could send the detectors off the charts if the instrument is not properly isolated. The trap is fully automated and remotely operated with custom-written LabVIEW software. The position of our trap in the sample image plane is controlled by a pair of acousto-optical deflectors and can be updated every 50 microseconds, which together with the computational speed of a Xilinx FPGA enables us to implement fast force feedback algorithms and perform accurate position detector calibrations. Among other experiments, we use the trap to apply forces to individual dynein motors walking along axonemes, allowing us detailed insight into the stepping behavior and mechanochemistry of the motor. Additionally, we have two TIRF lines coupled into our setup (488nm and 633nm) so that we have the option to observe single fluorophores on a highly sensitive Andor EMCCD camera.
Tracking the fate of single fluorescently-labeled proteins moving in live cells for long periods is often complicated because of the proteins' high density. Even if single particles are briefly visible, their paths cross and become indistinguishable. To address this, our lab developed the Photogate, a method for visualizing single-particle dynamics in crowded environments. We use a focused laser beam to photobleach the fluorescent molecules in an area, then briefly allow the fluorescence to recover from outside of the bleached area. After a few molecules enter the region of interest, we periodically bleach the periphery to prevent further fluorescent molecules from entering unbleached. This allows us to unambiguously observe the few fluorescent molecules inside the "gate." Using this method, we have visualized the dynamics of intraflagellar transport train turnaround at the ciliary tip, observed ligand-induced dimerization of a receptor tyrosine kinase at the cell surface, and directly measured binding and dissociation of signalling molecules from early endosomes in a dense cytoplasm, all at single-molecule resolution.
The ciliary tip is crowded with fluorescent signal from many IFT trains, but the Photogate allows us to see how proteins from single trains moving to the tip re-organize onto trains moving back towards the cell body.
A cell surface is heavily decorated with fluorescently labelled molecules (blue circles). The focused gate beam is swept outwards from the centre in a spiral pattern to pre-bleach an elliptical region. The gate beam is turned off to allow diffusion of fluorescent molecules from the periphery of the ROI, which are then imaged under TIRF illumination (yellow circles). The gate beam is repeatedly turned on to photobleach fluorescent particles entering the ROI (dark blue circles) while single molecules inside the ROI are imaged under TIRF illumination.