Ricardo R. Brau

Passive and active microrheology with optical tweezers

Efforts at understanding the behaviour of complex materials at the micro scale have led to the development of many microrheological techniques capable of probing viscoelastic behaviour. Among these, optical tweezers have been extensively developed for biophysical applications: they offer several advantages over traditional techniques, and can be employed in both passive and active microrheology. In this report, we outline several methods that can be used with optical tweezers to measure the microrheological behaviour of materials such as glycerol, methylcellulose solutions, actin matrices, and cellular membranes. In addition, we quantify the effect that the index of refraction of the solution has on the stiffness of the optical trap. Our results indicate that optical tweezers force microscopy is a versatile tool for the exploration of viscoelastic behaviour in a range of substrates at the micro scale.

Single M13 bacteriophage tethering and stretching

The ability to present biomolecules on the highly organized structure of M13 filamentous bacteriophage is a unique advantage. Where previously this viral template was shown to direct the orientation and nucleation of nanocrystals and materials, here we apply it in the context of single-molecule (SM) biophysics. Genetically engineered constructs were used to display different reactive species at each of the filament ends and along the major capsid, and the resulting hetero-functional particles were shown to consistently tether microscopic beads in solution. With this system, we report the development of a SM assay based on M13 bacteriophage. We also report the quantitative characterization of the biopolymers elasticity by using an optical trap with nanometer-scale position resolution. Expanding the fluctuating rod limit of the wormlike chain to incorporate enthalpic polymer stretching yielded a model capable of accurately capturing the full range of extensions. Fits of the force-extension measurements gave a mean persistence length of ≈1,265 nm, lending SM support for a shorter filamentous bacteriophage persistence length than previously thought. Furthermore, a predicted stretching modulus roughly two times that of dsDNA, coupled with the systems linkage versatility and load-bearing capability, makes the M13 template an attractive candidate for use in tethered bead architectures.

Detecting force-induced molecular transitions with fluorescence resonant energy transfer.

Single-molecule techniques have been responsible for substantial advances in the field of biophysics. Among these approaches, single-molecule fluorescence resonant energy transfer (FRET) spectroscopy provides an experimental view of the structural properties of individual molecules, whereas optical-tweezers force microscopy allows direct manipulation of the reaction coordinate of a single molecule. However, the simultaneous application of these techniques is complicated by optical-trap-induced photobleaching, which substantially reduces fluorophore longevity to unacceptably short time-scales. Herein, we describe a general solution to this problem and apply it to a novel force sensor based on a DNA hairpin, in the first successful combination of optical trapping and FRET. By alternately exposing the sample molecule to the optical-trapping and fluorescence-excitation lasers, we demonstrate the ability to reversibly manipulate a single molecule while simultaneously monitoring its structural configuration. This integrated measurement provides high-resolution mechanical control over molecular conformation with fluorescence-based structural reporting. The application of this technique for single-molecule exploration will lead to new experiments that employ combined optical trapping and single-molecule fluorescence for the simultaneous and active manipulation and monitoring of molecular structure in real time. Single-molecule force microscopy and fluorescence spectroscopy reveal individual molecular properties that are clouded by the inherent averaging of ensemble methods. However, the individual approaches of these techniques often fail to uncover the interplay between applied mechanical forces and structural changes. A single measurement of a force-sensing molecule connects these two perspectives by directly manipulating a molecular reaction coordinate while simultaneously detecting localized structural effects. Among the biophysical techniques capable of probing single-molecule properties, optical-tweezers force microscopy operates at piconewton force levels that are optimal for the detection of nanometer-scale conformational transitions. Likewise, single-molecule FRET spectroscopy provides complementary information about dynamic structural properties, including environment, orientation, and proximity, with comparable spatial resolution.[1] Previous efforts to combine these two techniques for a single, coincident measurement have been complicated by accelerated photobleaching rates induced by the high-intensity optical trap. Because of this effect, which is especially pronounced in common single-molecule FRET donor labels such as the dyes Cy3 and Alexa 555,[2] previous advances towards combining these techniques have spatially separated the fluorescent markers from the optical trap[3] or have employed uniquely robust chromophores.[4] We recently described a broadly applicable solution to this problem by alternately modulating the fluorescence-excitation and optical-trapping beams, which dramatically reduced this phenomenon without compromising trap integrity.[5] Herein, we show that such an optical modulation can be adapted to extend the emission times of FRET-paired labels without otherwise affecting their photophysical properties. To demonstrate this technique, we describe the first combination of optical-tweezers force microscopy with the single-molecule FRET detection of a novel force-sensing molecule into a single, integrated method capable of actively controlling molecular structure while simultaneously monitoring the conformational state of a single DNA hairpin molecule. The mechanics of DNA hairpins have been studied at the single-molecule level and, thus, offer a benchmark for examining optical tweezers and single-molecule FRET in a combined arrangement. These structures, which are commonly used to model secondary structure in nucleotides, are readily adapted for the mechanical exploration of conformational dynamics, as they undergo a sequence-dependent, reversible unzipping transition.[6,7] In addition, alternate constructs have been adapted for force-sensing applications.[8] The structure used in this work, which contains a 20-base-pair hairpin stem, is flanked by noncomplimentary sequences annealed to oligonucleotides functionalized with the fluorophores Cy3 and Alexa 647 (Figure 1). Complexes exhibiting single-molecule FRET emission were mechanically loaded with the optical trap, effectively reducing the energetic barrier to hairpin opening. This unzipping transition, which occurs at a force of approximately 18 pN, comparable to other similar measurements,[7] was reflected by the displacement of the bead toward the center of the trap. The conformational transition was accompanied by a simultaneous reduction in FRET efficiency caused by the increased physical separation of the Cy3 donor and the Alexa 647 acceptor, which indicated the precise location of the structural change caused by the translation of the mechanical load between the low-force (ca. 6 pN) and high-force (ca. 24 pN) states (Figure 2). The DNA complexes were moved through several transitions in a process corresponding to the reversible opening and closing of the hairpin segment, which demonstrated both the high degree of mechanical control and the simultaneous reporting by FRET emission. Furthermore, in the representative trace, single-step photobleaching of the donor after approximately 65 s verified the single-molecule measurement. Figure 1 Experimental assay design (see Experimental Section for details). DNA hairpin complexes, labeled with opposing Cy3 and Alexa 647 fluorophores, were mechanically loaded by translating the coverslip, as the position of the trapped bead and the emission ... Figure 2 Mechanically induced conformational changes monitored with FRET spectroscopy. A) A DNA hairpin was manipulated with optical tweezers between open or closed conformational states (black) that transition at loads of approximately 18 pN. The state of the ... This combination of optical-tweezers force microscopy and single-molecule FRET detection represents a significant advance for measuring the effects of structural changes on molecular function in a single molecule. By mechanically altering the conformational energy landscape, we actively induced a structural rearrangement pinpointed by strategically placed fluorescence labels. With minor modifications to existing assays, this approach can be extended beyond this model system to provide important new insight into the localized effects of mechanical force in biomolecular systems. For example, this combined technique can be adapted to monitor the intermolecular processes involved in the formation of a mechanically loaded protein complex,[9] the effects of mechanical deformation on single-enzyme catalysis,[10] or the intramolecular movements involved in biological-motor motility.[11,12] In addition, the presence of quantized single-molecule fluorescence signals can provide unambiguous verification of the size and location of a mechanical event, a critical tool for the design of often complex single-molecule assays. The new perspective that arises from this ability to physically deform single molecules while simultaneously measuring structural changes will allow the design of novel force-sensing molecules and will permit a new class of experiments for probing the interrelationship between molecular structure and biochemical function.

Single-molecule denaturation and degradation of proteins by the AAA+ ClpXP protease

ClpXP is an ATP-fueled molecular machine that unfolds and degrades target proteins. ClpX, an AAA+ enzyme, recognizes specific proteins, and then uses cycles of ATP hydrolysis to denature any native structure and to translocate the unfolded polypeptide into ClpP for degradation. Here, we develop and apply single-molecule fluorescence assays to probe the kinetics of protein denaturation and degradation by ClpXP. These assays employ a single-chain variant of the ClpX hexamer, linked via a single biotin to a streptavidin-coated surface, and fusion substrates with an N-terminal fluorophore and a C-terminal GFP-titin-ssrA module. In the presence of adenosine 5′-[γ-thio]triphosphate (ATPγS), ClpXP degrades the titin-ssrA portion of these substrates but stalls when it encounters GFP. Exchange into ATP then allows synchronous resumption of denaturation and degradation of GFP and any downstream domains. GFP unfolding can be monitored directly, because intrinsic fluorescence is quenched by denaturation. The time required for complete degradation coincides with loss of the substrate fluorophore from the protease complex. Fitting single-molecule data for a set of related substrates provides time constants for ClpX unfolding, translocation, and a terminal step that may involve product release. Comparison of these single-molecule results with kinetics measured in bulk solution indicates similar levels of microscopic and macroscopic ClpXP activity. These results support a stochastic engagement/unfolding mechanism that ultimately results in highly processive degradation and set the stage for more detailed single-molecule studies of machine function.

Excitation nonlinearities in emission reabsorption laser-induced fluorescence techniques

The effects of the nonlinear behavior of fluorescent intensity with excitation intensity on emission reabsorption laser-induced fluorescence (ERLIF) are investigated. Excitation nonlinearities arise mainly as a consequence of the depletion of the ground-state population stemming from the finite lifetime of molecules in the excited state. These nonlinearities hinder proper suppression of the excitation intensity information in the fluorescence ratio, degrading measurement accuracy. A method for minimizing this effect is presented. This method is based on the approximation of the fluorescence intensity nonlinearities by a power law. Elevating the two-dimensional fluorescent intensity maps to the appropriate exponent allows for proper suppression of excitation intensity in the fluorescence ratio. An overview of the principles and constitutive equations behind ERLIF film-thickness measurements, along with a characterization of the fluorescences nonlinear behavior, is presented. The power law approximation and processing scheme used to mitigate this behavior are introduced. Experimental proof of the validity of the approximation and processing scheme is provided.

IOFF generally extends fluorophore longevity in the presence of an optical trap.

The combination of optical tweezers force microscopy and single molecule fluorescence has previously been complicated by trap-induced photobleaching. Recent studies have suggested that this effect is caused by a sequential absorption of photons, leading to ionization of the fluorescent singlet state. In this work, we show the range of effects of optical trapping radiation on common fluorescent dyes. Using the interlaced optical force fluorescence (IOFF) laser modulation technique, we show that the removal of simultaneous near infrared radiation dramatically reduces photobleaching effects. However, these studies show that the sequential addition of near infrared radiation in some cases extends photobleaching longevity beyond the natural intrinsic decay. We suggest a refined photoelectronic mechanism that accounts for the possibility of reverse intersystem crossing from a reactive triplet state and explains the nature of trap-induced photobleaching.

Clpxp Degradation of Proteins Probed By Single-Molecule Fluorescence

ClpXP is an AAA+ protease that unfolds and degrades target proteins. ClpX, a hexameric ring-shaped ATPase, recognizes specific proteins and then powers their mechanical denaturation and translocation into the degradation chamber of ClpP where polypeptide bond cleavage occurs. Although ClpXP degradation activities have been widely studied at the bulk solution level, the operating principles and detailed mechanisms of this complex macromolecular machinery remain unanswered. Here, we probe the kinetics of substrate unfolding and degradation by ClpXP using a single-molecule fluorescence assay. These assays employ a covalently crosslinked ClpX hexamer, immobilized on PEG coated surface illuminated by total internal reflection fluorescence. A series of substrates are engineered to contain fusion of an N-terminal Cy3 and a C-terminal GFP-titin-ssrA module. In the presence of ATPγS, ClpX stalls at GFP after degradation of titin-ssrA domains. These stalled pre-engaged substrates are stably bound to ClpXP even in the absence of Mg++, but are released quickly upon the introduction of nucleotide-free solution. Exchange into ATP for pre-engaged substrate-ClpXP complexes allows synchronous resumption of unfolding and degradation of GFP and any following domains. The time required for complete degradation is measured by loss of the N-terminal Cy3 from the protease complex. GFP unfolding can also be monitored directly with quenching of intrinsic fluorescence by denaturation. Global fitting of single-molecule data for a set of related substrates yields time constants for ClpX unfolding, translocation, and a terminal step which may involve product release, and shows strong agreement with bulk solution measurements. It should be possible to extend these methods to allow single-molecule studies such as FRET for real-time assays of ATP-fueled conformational changes that drive the mechanical operations of the ClpXP protease. Support from the NSF Career Award (0643745) is gratefully acknowledged.