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Week #
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Topics
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Lecture Summaries
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1
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Introduction to Single Molecule Imaging Techniques
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This first class will be an introduction to the course, students, and the instructor. We will begin by introducing ourselves and our backgrounds and interests. We will go over the syllabus, goals and expectations for the course and schedule meeting times. The instructor will introduce the core concepts of the microscopy techniques used in single-molecule imaging, discuss the rapid development of these methods the last two decades, and describe some key applications to studies of DNA replication, transcription, and cytoskeletal structure. This introduction will provide an overview of the techniques as well as a brief introduction to the subjects we will focus on for the rest of the semester.
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2–4 Part One: Manipulation of Polymers by Force
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2
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Single Optical Traps
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Optical traps, also known as optical tweezers, provide unique means to control the dynamics of small particles. It is possible to remotely apply forces on living cells, internal parts of cells and large biological molecules within cells. One of the most important applications is the study of motor molecules, such as kinesin and myosin. With optical traps, which use a highly focused laser beam to hold and move microscopic objects, one can measure the forces generated by single motor molecules and resolve their detailed stepping motion.
To explore the different types of optical traps and their uses in combination with other techniques, we will start with single optical traps. The first paper, Block et al., studies the kinetics of a kinesin motor molecule. In a more recent study, Sheinin et al., a different kind of single optical trap, which is called angular optical trapping technique that applies torque and force simultaneously to a trapped particle, is developed for single-molecule torsional studies of DNA and its processing machineries, also called rotary motors.
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3
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Magnetic tweezers
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Magnetic force spectroscopy is a rapidly developing single-molecule technique that has found numerous applications at the interface between physics and biology. Since the invention of the first magnetic tweezers or traps, where similar to optical traps, a magnetic bead (hence anything attached to the magnetic bead) is held and manipulated within a magnetic field, several modifications to the approach have helped to relieve the limitations of the original design while amplifying its strengths.
In session 2, we learned that the main advantage of optical tweezers is the ease with which single beads can be manipulated and carried around. In contrast, magnetic tweezers are at their best when working with a homogeneous force field and manipulating multiple beads at once. In Dulin et al. a magnetic trap is used to monitor the opening and closing of a DNA hairpin on millisecond timescales in real time. In van der Heijen et al. magnetic tweezers are used to study the dynamics of assembly of a RecA–DNA filament (RecA is a protein that covers exposed ssDNA during repair) on a torsionally constrained DNA molecule in real time.
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4
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Double optical traps
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In dual-trap systems, the molecule is tethered between two optically trapped beads. The choice of single-trap versus dual-trap design is often dictated by the system to be studied. Surface-based applications, meaning that at least one end of a molecule is attached to a surface, as in single optical traps where DNA molecule is attached to the surface at one end, and the other end is attached to bead which is held above the surface with a laser beam, are subject to unwanted motion of the sample stage or ‘drift,’ which can be significant and limit measurement resolution. In dual-trap set-ups the traps are decoupled from the surface and its drift.
To explore other advantages and disadvantages of dual traps, we will discuss Lisica et al., an exciting application of dual optical traps in which RNA polymerase is monitored as it moves along a DNA molecule and transcribes an RNA strand. The trajectories (velocity, pausing, backtracking, etc.) of the RNA polymerases PolI and PolII on DNA are monitored in real time and analyzed in detail using dual trap setups. Gardini et al. describe a 3-bead version of dual optical traps to study kinetics of the motor-protein myosin.
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5–7 Part Two: Fluorescence Microscopy
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5
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Total Internal Reflection Fluorescence Microscopy (TIRFM)
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Fluorescence-based single-molecule techniques allow the real-time observation of the trajectory of molecules labeled with single fluorophores, which are excited with a laser of the appropriate wavelength. There are two complementary fluorescence techniques that differ in their excitation and detection modalities, total internal reflection fluorescence (TIRF) and confocal (confocal microscopy will be discussed in Session 9). TIRFM is a powerful technique for selectively imaging fluorescent molecules (e.g., GFP, membrane dyes, fluorochromes attached to antibodies, etc.) in an aqueous environment that are very near a solid substance with a high refractive index (e.g. coverslip). While the thickness of a confocal image section is approximately 500 nm, it is only 100 nm in TIRF.
The advantages and disadvantages of TIRF will be discussed as we consider Yardimci et al., who visualize the uncoupling of sister replisomes using TIRF microscopy and frog egg extracts. In Thawani et al. TIRF microscopy is used to visualize nucleation of microtubules and reveal kinetics in the transition states.
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6
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Single-Molecule FRET (smFRET)
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Single-molecule fluorescence resonance energy transfer (smFRET) is one of the most general and adaptable single-molecule techniques. Since its humble beginning under non-aqueous conditions in 1996, smFRET has rapidly developed to answer fundamental questions about replication, recombination, transcription, translation, RNA folding and catalysis, non-canonical DNA dynamics, protein folding and conformational changes, various motor proteins, membrane fusion proteins, ion channels, and signal transduction, to name just a few, and the list keeps growing at a fast pace. In FRET measurements, the extent of non-radiative energy transfer between two fluorescent dye molecules—termed donor and acceptor—reports the intervening distance, which can be estimated from the ratio of acceptor intensity to total emission intensity. Conformational dynamics of single molecules can be observed in real time by tracking FRET changes.
In Duchi et al., an optimized smFRET strategy is used to monitor de novo RNA synthesis in real time by monitoring DNA scrunching*. In Ticau et al., smFRET also reveals the intrinsic details of the sequential loading events during DNA replication initiation.
* RNA polymerase remains stationary while it unwinds and pulls downstream DNA into the transcription complex to pass the nucleotides through the polymerase active site, thereby transcribing the DNA without moving. This causes the unwound DNA to accumulate within the enzyme, hence the name “DNA scrunching."
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7
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DNA Curtains and Extensions by Flow
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ssDNA (single-strand DNA), dsDNA (double-strand DNA), and protein/nucleosome-bound DNA behave different under force. In 1953, the double-helical structure of DNA was described in a landmark paper by James Watson and Francis Crick. That paper was a game changer, as it stated that DNA has two helical chains each coiled round the same axis and has the bases on the inside and the phosphates on the outside of the helix. More importantly, the novelty of the structure reflected the way in which the two chains are held together by the purine and pyrimidine bases: “They are joined together in pairs, a single base from one chain being hydrogen-bonded to a single base from the other chain so that the two lie side with identical z-coordinates.” This description of the structure of DNA was just the beginning, and our knowledge of DNA has increased exponentially after this crucial study. Biological, physical, chemical and mechanical properties of DNA have been analyzed extensively.
In Kim et al., DNA combing, where DNA molecules are stretched horizontally and attached from two ends in a microfluidic chamber (which makes DNA molecules look like “combed” hair on a surface) and TIRF microscopy, which is a fluorescence microscopy technique that only excites molecules in a limited specimen region and hence greatly reduces background signal (as discussed in Session 5), reveal how much force DNA molecules are under during transcription. In Kong et al., shortening of DNA upon compaction is utilized to understand mechanical properties of condensins, enzymes that are considered the primary driver of chromosome architecture.
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8–10 Part Three: Optical Trap and Fluorescence Meet-up
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8
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Fleezers: Combining Fluorescence with Optical Tweezers
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Hybrid instruments integrate standard optical trap layouts with fluorescence excitation lasers and detection optics. Hybrid instruments combining fluorescence with optical tweezers (fleezers) have become increasingly popular, as they enable measurements of mechanical signals and forces simultaneously with fluorescence imaging. A wide variety of fluorescence-trap instrument designs have been implemented integrating both single-trap and dual-trap configurations with wide-field epifluorescence, total internal reflection (TIRF) and confocal microscopy1.
We will discuss a powerful application of single trap fleezers in Hilario et al., where they visualize human Rad512 assembly and disassembly on duplex DNA, and dual trap fleezers in van Mameren et al., in which the molecular mechanism of RAD51 filament disassembly is unraveled.
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9
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Fleezers 2.0-Dual Traps Combined with Confocal Microscopy
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In confocal microscopy, the excitation light is instead focused by the objective to a diffraction-limited spot (~250 nm in diameter) inside the sample, and light emitted only from this spot is collected. When combined with optical traps, this instrument has the ability to resolve mechanical signals at subnanometer spatial resolution (with the optical traps) and to detect simultaneously the emitted light from a single fluorophore (with the confocal microscope). The ability to measure and/or control multiple variables simultaneously has provided many new insights not possible when using each method independently.
Desai et al. use this application to study translation regulation and Wasserman et al. analyze the activation of the eukaryotic helicase CMG by constraining duplex DNA between two traps and monitoring fluorescently labelled proteins with confocal microscopy.
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10
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Optical Traps Meet both Confocal and FRET
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As FRET measures the distance separating a dye pair, it can be combined with optical trapping to monitor, as a function of force, distance changes caused by conformational interconversions of nucleic acid structures and nucleoprotein complexes.
We will discuss Ngo et al. to understand how they utilized this approach to study nucleosome dynamics. As FRET provides measurements of inter-dye distances independent of those obtained from the optical traps, it can be used to track reaction coordinates to which the traps are insensitive. The access to ‘orthogonal’ reaction coordinates enables mapping of multidimensional energy landscapes.
Suksombat et al. is a beautiful example of this application – similar to Ngo et al., they applied it to understand the wrapping topologies of the ssDNA binding protein SSB.
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11–13 Part Four: Live Cell Imaging Applications
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11
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Stochastic Optical Reconstruction Microscopy (STORM)
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In 2014, the Nobel Prize in Chemistry was awarded jointly to Eric Betzig, Stefan W. Hell and William E. Moerner “for the development of super-resolved (super-res) fluorescence microscopy.” Super- resolution microscopy, or nanoscopy, provides the ability to see details of cellular and even macromolecular structure that were not possible to see before. Notably, nanoscopy is compatible with live cells and has the capability for multiplex labeling with high molecular specificity.
In Xu et al., super-res microscopy is used to reveal fine structural details of axons. Balint et al. use super-res microscopy for live-cell imaging to study cargo dynamics at microtubule intersections.
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12
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Photoactivation Localization Microscopy (PALM) and Stimulated Emission/Depletion (STED)
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The development of photoactivation localization microscopy (PALM) is closely linked to the advent of photoactivatable proteins, which allow us to use light to control the density of fluorescing proteins in each image. PALM takes advantage of the versatility and specificity of genetically encoded, fluorescently tagged molecules in cells, and has quickly become the tool of choice for super-resolution live-cell imaging.
The stimulated emission depletion (STED) microscope, on the other hand, provides spatial resolution well below the limit imposed by the diffraction of light. The STED microscope was invented in 1994, but has gained substantial momentum only in the last 10 years.
Rossier et al. use single-protein tracking and super-resolution imaging to visualize the dynamic nano-organizations of integrins and talin inside focal adhesions (FAs)*. In Hein et al., STED is used to demonstrate far-field optical imaging with subdiffraction resolution of the endoplasmic reticulum (ER) in the interior of a living mammalian cell.
*Focal adhesions (FAs): Focal adhesion is a type of adhesive contact between the cell and extracellular matrix through the interaction of the transmembrane proteins integrins with their extracellular ligands, and intracellular multiprotein assemblies connected to the actin cytoskeleton.
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13
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Structured Illumination Microscopy (SIM) and Single-Molecule Localization Microscopy (SMLM)
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Structured illumination microscopy (SIM) is based on standard wide-field microscopy and is compatible with most standard fluorophores and labeling protocols. SIM is an attractive choice for live-cell imaging; it requires no special fluorophores or high light intensities to achieve twice diffraction-limited resolution in three dimensions. SIM provides an approximately twofold resolution enhancement of standard wide-field microscopy as compared to other super-resolution methods.
Fiolka et al. use SIM to record 3D two-color datasets of living whole cells. For the rest of the class, we will discuss students’ prospective papers that they plan to present in the final week as their final assignment.
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14
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Oral Presentations and final discussions
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Students will give oral presentations (~ 20 min) about their chosen research papers and lead critical discussions about the papers’ experimental design, data and conclusions. We will have a final discussion about what we have learned about single-molecule techniques, their applications in different fields, the challenges of the field and future directions. We will discuss what worked well and what did not in the course. Finally, students will complete written course evaluations and provide additional feedback to the instructor about the course.
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