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Integrating Optical Tweezers, DNA Tightropes, and Single-Molecule Fluorescence Imaging
CHAPTER SEVEN
Integrating Optical Tweezers, DNA Tightropes, and Single-Molecule Fluorescence Imaging: Pitfalls and Traps J. Wang*, J.T. Barnett*, M.R. Pollard†, N.M. Kad*,1 *School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom † DFM A/S, Kongens Lyngby, Denmark 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Elongating Bundled DNA for Imaging 2.1 DNA Combing 2.2 DNA Tightropes 3. Integrating Laser Tweezers Into Biological Experiments 3.1 Introduction 3.2 Trapping With Nanoprobes 4. Controlling and Detecting the Nanoprobe 4.1 Using an AOD to Create Three Traps 4.2 Detection Strategies 5. Applying the Nanoprobe to Biological Study Systems 5.1 Measuring the Tension of a DNA Tightrope 5.2 Measuring Interactions With Single Proteins 6. Conclusions and Outlook References
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Abstract Fluorescence imaging is one of the cornerstone techniques for understanding how single molecules search for their targets on DNA. By tagging individual proteins, it is possible to track their position with high accuracy. However, to understand how proteins search for targets, it is necessary to elongate the DNA to avoid protein localization ambiguities. Such structures known as “DNA tightropes” are tremendously powerful for imaging target location; however, they lack information about how force and load affect protein behavior. The use of optically trapped microstructures offers the means to apply and measure force effects. Here we describe a system that we recently developed to enable individual proteins to be directly manipulated on DNA tightropes. Proteins bound to DNA can be conjugated with Qdot fluorophores for visualization and also Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.003
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2017 Elsevier Inc. All rights reserved.
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directly manipulated by an optically trapped, manufactured microstructure. Together this offers a new approach to understanding the physical environment of molecules, and the combination with DNA tightropes presents opportunities to study complex biological phenomena.
1. INTRODUCTION Proteins and substrates exist in a physical milieu where they are subject to processes such as Brownian motion, diffusion, and external force. The contribution of these factors toward the overall mechanism requires the properties of the individual components to be determined for extrapolation to the ensemble system. One major biological process that requires understanding at this level is diffusion to target, and particularly so for targets on DNA. Based on diffusion models, it is possible to predict the association time of a protein to its target site (Berg, 1993). However, Lac repressor was found to locate its target site faster than predicted by a three-dimensional (3D) encounter model (Riggs, Suzuki, & Bourgeois, 1970). Facilitated diffusion was postulated to explain this discrepancy and involves a change in the mechanism of target search from being wholly 3D to also include onedimensional (1D) diffusion along the backbone of the DNA (Adam & Delbr€ uck, 1968; Winter, Berg, & von Hippel, 1981). For many years, numerous investigators set out to test this hypothesis (Halford & Marko, 2004) prior to the application of relatively new single-molecule methods. These methods permit the observer to track a single molecule labeled with a fluorophore as it moves toward its target site. Early investigations proved the existence of 1D sliding (Blainey, van Oijen, Banerjee, Verdine, & Xie, 2006; Tafvizi et al., 2008), hopping (Bonnet et al., 2008) which is longdistance jumps in position along a single DNA strand and also a combination of all of these mechanisms (Hughes et al., 2013; Kad, Wang, Kennedy, Warshaw, & van Houten, 2010). Also clear was that some proteins were capable of directional motor movement (van Oijen et al., 2003). To study such systems at the single-molecule level, first it requires DNA to be elongated. In Fig. 1 we show an image of a field of λ-DNA molecules stained with YOYO-1 dye. λ-DNA (the genome of bacteriophage lambda) consists of 48,502 base pairs; with a base pair separation of 0.34 nm, this equates to a contour length of 16.5 μm. It is apparent in Fig. 1 that the molecules in the image have collapsed into a bundled conformation. The radius pffiffiffiffiffiffiffiffi of a bundle can be calculated using random flight theory r ¼ l K ,
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Fig. 1 Image of YOYO-1 stained bacteriophage λ-DNA molecules. This DNA has a contour length of 16 μm; however, in solution, the DNA collapses into bundles. These diffuse through the field of view and are also seen to change shape.
where l is the contour length and K is the Kuhn length (twice the persistence length), for λ-DNA this is 1.3 μm. Comparing this to the point spread function of a typical fluorophore (Anderson, Georgiou, Morrison, Stevenson, & Cherry, 1992; Thompson, Larson, & Webb, 2002), it is apparent that measuring motion within this bundle is extremely challenging. Furthermore, the bundle will move, so the positional accuracy is adversely affected. Here we describe how we construct DNA tightropes to achieve elongated DNA structures and how this can be combined with optically trapped probes for more sophisticated experiments.
2. ELONGATING BUNDLED DNA FOR IMAGING 2.1 DNA Combing DNA can be laid onto an activated surface (Allemand, Bensimon, Jullien, Bensimon, & Croquette, 1997), usually polystyrene or polymethylacrylate by a method known as combing (Bensimon et al., 1994). This is achieved by either retracting an activated slide through a solution of DNA or flowing DNA across a surface (Crut et al., 2005; Deen et al., 2015; Lyon, Fang, Haskins, & Nie, 1998). Typically performed at low pH to achieve better binding, the DNA can then be examined using total internal reflection fluorescence microscopy. This reduces the amount of background fluorescence permitting fluorescently labeled proteins on DNA to be clearly imaged. In addition to this approach the DNA can be attached by one end and flow applied to elongate the DNA such as used to study the DNA repair protein
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human oxoguanine DNA glycosylase 1 (Blainey et al., 2006) or DNA replication (Lee et al., 2006). Although more complex, this approach reduces, but does not eliminate, interactions between DNA-bound proteins and the surface. Visually, however, it is unclear whether the proteins are attached to the DNA or surface because both are in the same focal plane.
2.2 DNA Tightropes This method exploits the simplicity of combing but raises the DNA from the surface without the need for flow. Suspending individual DNA molecules between surface-immobilized beads ensures that anything visible within the focal plane above the surface must be bound to DNA and not the surface. To facilitate imaging, it is necessary to optimize illumination such that only the tightropes are excited and the rest of the solution receives minimal light. This is achieved by using oblique angle fluorescence excitation microscopy (Hughes et al., 2013; Konopka & Bednarek, 2008; Tokunaga, Imamoto, & Sakata-Sogawa, 2008). To create DNA tightropes we use the following protocol: 1. Plasma clean 22 40 mm coverslips (Harrick PDC-32G, Ithaca, NY) for 2 min per slide. 2. Build a flow chamber as described in Fig. 2. 3. Block the chamber overnight by flowing in mPEG5000 (Sigma Aldrich) pH 8.2, wash with water, and then block overnight again with 50 mM Tris–HCl, 10 mg/mL BSA, 0.1% Tween20. 4. Flow in 5 μm silica beads (precoated overnight with 350 μg/mL poly-Llysine (Sigma Aldrich)), until bead separation is on average 20 μm. 5. Wash flow chamber with 200 μL buffer and check bead density again to ensure that beads have not been washed from the surface. In that event recoat beads with poly-L-lysine before adding to flow chamber (see Fig. 2 legend for details on constructing a flow chamber; also see Kad et al., 2010). 6. Attach a syringe pump (AL-1000 World precision instruments) to one flow chamber outlet and to the other a microcentrifuge tube using polyethylene tubing (PE90; BD Medical, Oxford, UK). 7. Backflow the buffer from the syringe into the microcentrifuge tube to ensure an airtight connection. 8. Flow in 100 μL of 15 nM λ-DNA, this will bring the DNA into the flow chamber. Apply forward and backward flow to run the DNA across the surface beads for at least 20 min.
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Fig. 2 Design of flow cells for creating tightropes. To create flow chambers a plasma cleaned glass coverslip (#1.5) is attached to a glass microscope slide using an in-house cut double-sided adhesive gasket (UK industrial tapes) of depth 180 μm. The glass slide is predrilled (using a diamond bur; Precision Dental, London, UK) to allow the attachment of small diameter tubing and sealed using UV curable adhesive (NOA68, Thorlabs, Ely, UK) after insertion. The gasket design is shown in the lower panel, indicating the positions of ports used for tightropes and for nanoprobe injection. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
9. Flush the flow chamber with 200 μL of imaging buffer and if required 100 μL of 1 nM YOYO-1 dye to visualize the DNA. During experiments, it is usual to exclude the addition of YOYO-1 dye since it is known that anything in focus at the height of the beads will be bound to DNA.
3. INTEGRATING LASER TWEEZERS INTO BIOLOGICAL EXPERIMENTS 3.1 Introduction Optical trapping, also known as laser tweezing, permits physical manipulation of single molecules. This approach requires a high-energy laser beam focused into a fluid sample to interact with particles of higher refractive index than their surrounding solution. As a consequence the particles, typically polystyrene or silica beads, become fixed near the focal point of the laser
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beam. Deviation from the focal point results in a restoring force that is proportional to the distance from the center of the trap position over a region of approximately 500 nm. This Hookean response results in a very simple linear correlation between displacement and force. Therefore, not only are laser tweezers useful for physical placement of molecules but also their response to force. The application of laser tweezers for studying molecular motors was initially established with kinesin (Svoboda, Schmidt, Schnapp, & Block, 1993) and myosin (Finer, Simmons, & Spudich, 1994; Guilford et al., 1997; Molloy, Burns, Kendrick-Jones, Tregear, & White, 1995). The latter experiments elegantly suspend individual actin filaments between optically trapped beads, pull them taut (Dupuis, Guilford, Wu, & Warshaw, 1997), and then use this platform to investigate the properties of single myosin molecules attached to a solid surface (Fig. 3). These experiments have revolutionized biology by permitting the load response of single molecules to be determined (Kad, Patlak, Fagnant, Trybus, & Warshaw, 2007; Kad, Trybus, & Warshaw, 2008; Veigel, Molloy, Schmitz, & Kendrick-Jones,
Fig. 3 The three-bead assay. This assay, used primarily for studies of the molecular motor myosin, involves plating a low density of myosin onto a coverslip surface that is precoated with 3-μm silica beads. 1-μm silica beads are trapped and used to capture the ends of an actin filament, which is then tautened. When the actin is brought into contact with the surface-bound myosin and the position of the trapped beads detected, a clear drop in variance is seen. This is due to the additional contribution of the stage to the stiffness of the system. In addition, myosin will move the actin which is seen as a deflection from the baseline of the bead position.
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2003; Veigel, Schmitz, Wang, & Sellers, 2005). Such measurements are not possible at the bulk level and also permit the contribution of the single molecule in the context of the ensemble to be determined. Such scaling measurements are integral to the construction of nanobiotechnological devices (Hess, 2011) and for the understanding of complex biological systems. Beyond molecular motors the study of DNA-based enzymes has also been facilitated by the use of laser tweezers. This has typically taken the form of suspending single DNA molecules between beads in much the same way as actin has been used for studying myosin (Hilario, Amitani, Baskin, & Kowalczykowski, 2009; Qi, Pugh, Spies, & Chemla, 2013; Skinner, Baumann, Quinn, Molloy, & Hoggett, 2004; Wang, Yin, Landick, Gelles, & Block, 1997). However, recently more elaborate applications of laser tweezers have been used to study the localization and affinity of proteins for DNA. These include looping one DNA molecule around another (Noom, van den Broek, van Mameren, & Wuite, 2007; van Loenhout et al., 2013) or incorporating atomic force microscope probes (Huisstede, Subramaniam, & Bennink, 2007; Shon & Cohen, 2016).
3.2 Trapping With Nanoprobes 3.2.1 Fabrication and Preparation of Nanoprobes Optical trapping has traditionally involved the use of spherical beads that are isotropically suspended in the traps. This means that the movement of the beads is limited to simple translation and that the surface available for interacting with molecules is large and cannot be easily controlled. To overcome these limitations, we and others have used structured particles that can be optically trapped. As shown in Fig. 4 we use a triangular structure with three
Fig. 4 The structure of the nanoprobe. Fabricated using EBL from SU8 the nanoprobe is a triangular structure with three cylinders located at each vertex. Three laser traps are used to position the probe each focused on a different cylinder. The probe is a long thin blade structure, and the nanoprobe is visible by fluorescence as well as bright-field microscopy. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., Botchway, S. W., Parker, A. W., & Kad, N. M. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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trapping points and a protruding tip (Pollard et al., 2010). This permits accurate 3D positioning of the protrusion such that molecules can be manipulated with greater precision. The “nanoprobe” structures were fabricated using electron beam lithography as follows. To etch the nanoprobes a chrome layer followed by an SU8 photoresist layer is applied to a silicon wafer, baked, and then exposed (Pollard et al., 2010). This process yielded 4000 nanoprobes per segment on a silicon wafer. Correct release of the nanoprobes from the wafer is pivotal to enable subsequent experiments to be performed efficiently. First, the chrome layer is dissolved using ceric ammonium nitrate (CAN) pipetted directly onto the nanoprobe containing segment. The duration of exposure to etchant is controlled in order to leave some chrome connecting the nanoprobes to the wafer. This permits subsequent washing steps without loss of nanoprobes; any remaining CAN residue in the wafer region containing nanoprobes is gently washed away using high-purity water several times. Residual CAN was found to inhibit downstream steps, and washing also removes any free particulates that can contaminate the flow chamber and interfere with trapping experiments. After several washes the nanoprobes can be released with a more vigorous rinse with buffer containing BSA and Tween20 (Simons et al., 2015). This step prevents aggregation of the nanoprobes which are then collected and kept for use in the flow cell. 3.2.2 Flow Chamber Adaptations for Improved Experimental Yield For introduction of the nanoprobes into the flow chamber, it is necessary to avoid contact with the walls of the tubing, which was found to bind avidly to the nanoprobes. Therefore to add nanoprobes, a separate hole was drilled into the flow chamber requiring a specific design with a separate channel for nanoprobes (Fig. 2). This design not only avoids the use of polyethylene tubing during addition of nanoprobes but also creates a channel for storage of nanoprobes within the flow chamber. Nanoprobes are injected using a glass Hamilton syringe, and the channel can be sealed simply using commercially available sticky tape.
4. CONTROLLING AND DETECTING THE NANOPROBE Laser tweezers are relatively simple to construct, requiring only the back aperture of a high numerical aperture (>1.2) objective lens to be slightly overfilled with a stable laser. To obtain the correct beam size requires
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the use of one or more telescopes in the optical path. Controlling the position of the laser tweezers can be achieved by a number of schemes; however, they all share a common point that the turning mechanism is conjugated with the back focal point of the objective lens. This ensures a stable wide region for trapping, for a much more extensive discussion on the tweezer setup used (see Sung, Sivaramakrishnan, Dunn, & Spudich, 2010). Later we discuss the control mechanism used for our system (Simons et al., 2015) and the detection scheme.
4.1 Using an AOD to Create Three Traps In order to manipulate the laser-trapping beam laterally across the imaging plane, acoustic optical deflectors (DTD-274HD6M, IntraAction) are inserted into the laser beam path to deflect the beams with varying acoustic frequency (Molloy, 1998). In such a way, three optical traps can be created to coincide spatially with the vertex positions of the nanoprobe. These are generated by rapidly (40 kHz) repositioning the laser between each position significantly faster than the nanoprobe response frequency of 30 Hz (Pollard et al., 2010). Removal of the IR-blocking filter permits an image of the trap position to be projected onto the camera, although this should only be performed when necessary to prevent damage to the camera. Once a floating nanoprobe is found in solution, adjust the orientation of the traps to match it, and note their position. Turn off the IR-trapping laser. Move the trap position marker over the nanoprobe and adjust the focus to be slightly above the nanoprobe. Turning on the IR laser should capture the nanoprobe. Since the nanoprobes are located in the side channel of the flow chamber (Fig. 2), they will need to be navigated to an appropriate DNA tightrope. This is a delicate operation requiring movements over millimeters, and touching anything in the flow chamber will likely result in the nanoprobe sticking. At high IR laser powers, it is possible to guide the nanoprobe over the surface beads. However, to image the tightropes simultaneously, the trapping and imaging planes need to be offset in the z-axis. This is achieved by moving the focusing lens for the optical tweezers. Once a tightrope is found, the lens is returned to its correct position bringing the fluorescence and trapping planes into parfocality. At this point, it is necessary to align the detection apparatus with the center of the nanoprobe vertices and to perform some calibrations of the trap stiffness. For the latter, position detection is required.
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4.2 Detection Strategies Classical detection of trapped objects requires either a position sensitive detector (PSD) or a quadrant photodiode (QPD). These record the position of the trapped object at the back focal plane of the microscope condenser. The recorded signal is an interference signal between the scattered light from the bead and incident trapping beam. This detection approach relies on the linear relationship between the center of mass of the interference pattern and the relative position between the trapping beam and the trapped bead; however, this scheme is not suitable for tracking multiple beads simultaneously. The output of a PSD, usually a voltage signal, will be affected by parameters such as power of the beam and its shape and size; therefore, careful calibration is required. We adopt a more flexible detection method of bead tracking by using image-based position detection which allows us to track multiple beads by following their movement. Previous studies (Belloni, Monneret, Monduc, & Scordia, 2008; Keen, Leach, Gibson, & Padgett, 2007; Otto, Gutsche, Kremer, & Keyser, 2008) have shown that image-based tracking can provide similar performance compared with PSD and QPD detection schemes with the added advantage of multiple trap detection with flexible controls. Multiple moving objects can be tracked using real-time updated region of interests (ROIs) across the image. With PSD or QPD detectors offering up to hundreds of kHz, image-based trapping cannot compete in terms of detection bandwidth. However, for most position measurements, a fast camera with a carefully positioned ROI will suffice. 4.2.1 Optics for Image-Based Detection The position detection path and the laser-trapping beam are built around a wide-field microscope. As shown in Fig. 5, a high-power white LED light source is placed above the sample stage of the microscope in a transmission configuration to provide bright-field illumination. A Hamamatsu ORCA-Flash 2.8 CMOS camera (Hamamatsu Photonics, Hamamatsu, Japan) with 1920 1440 pixels is used as the image sensor to acquire brightfield microscopic images. The imaging signal collected by the microscope objective, OBJ, is focused by a doublet achromatic lens (L1, f ¼ 200 mm) to form a conjugate imaging plane where a mechanical pupil (A) is placed to control the aperture size. A pair of achromatic lenses L2 (f ¼ 100 mm) and L3 (f ¼ 50) form a 4F configuration to relay the image at A to the image sensor. The trapping laser is a 5W 1070-nm ytterbium fiber-coupled laser that is separated from the detection path by a dichroic beam splitter. The laser beam is focused by the objective to the sample plane, SP, within the
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Fig. 5 Layout of the camera-based position detection scheme. CMOS, Hamamatsu’s Scientific CMOS Camera, ORCA-Flash 2.8; DBS, dichroic beam splitter; A, aperture; L1, L2, and L3, achromatic lens with focal length of 200, 100, and 50 mm, respectively; LED, brightfield light source; NBF, notch-blocking filter; OBJ, microscope objective; SP, sample plane; the dashed arrows indicate the positions of image plane and image conjugate planes.
field of view of the camera. A notch-blocking filter is used to stop the back reflection of the trapping beam reaching the camera. In this way, the camera can deliver the image of the SP with the trapping laser focus in the middle. Each pixel on the camera covers an area of 120 120 nm2 in the SP; therefore, an ROI of 16 16 pixels produces a detection window of 1.92 1.92 μm2. With this ROI, the ORCA-Flash 2.8 camera runs at a maximum frame rate of 1 kHz. The pixel size is calibrated by using a graticule (Graticules Ltd., Tonbridge, UK). To initially locate objects for trapping the full field of view is used. ROIs are then adopted to isolate individual objects floating in the solution. 4.2.2 Centroid Detection We describe here the use of 1.5-μm diameter silica beads as a trapped object; however, the procedure for a nanoprobe vertex is identical. When the bead is trapped by the laser, the image of the bead appears bright in the middle, providing an image with high contrast and small size. The center of mass is calculated by using the centroid approach in LabView™. N X N X
Cx ¼
i I ij
i¼1 j¼1 N X N X I ij i¼1 j¼1
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The centroid position in x (Cx) takes the sum of the product of the intensity of each pixel in a column (Iij) and that column position in x (i). This is repeated for every column in the ROI (i ¼ 1 to N). Finally the sum of all of these values is expressed relative to total intensity of the image. This calculation is repeated for the y-axis but for each row. We normally threshold the image to offset problems of low contrast and to remove any distractions from background features in the flow chamber. Using this detection system clear steps (<15 nm, pixel size ¼ 120 nm) in either the “x”- or the “y”-direction of a 1.5-μm bead are visible when the sample is translated by a nanometer-accuracy piezo stage (LPS200 Mad City Labs). Statistical treatment of the data can provide much higher resolution, and when trapped, further correlation methods can be applied (Mehta, Finer, & Spudich, 1997). 4.2.3 Acquisition and Processing Software Two pieces of software were created to control the camera and analyze the data from an acquired image sequence, both written in LabView™ (National Instruments, Newbury, UK). The camera control software sets up the parameters for the CMOS camera, acquires, and can export the image data to a hard drive when the sequence is too large for system RAM (using 32-bit Windows XP PC this is limited to 3 GB). Centroid information for the trapped bead is concurrently reported in real time. A raw image of a 1.5μm bead is displayed Fig. 6 (top), and the centroid determined “x” and “y” positions are plotted alongside. Using off-line software (Fig. 6, bottom), it is possible to generate image intensity histogram information, apply new thresholds for recalculation of centroid coordinates values, and visualize the processed bead position with a two-dimensional mapping. This validated detection scheme offers the capability of using more versatile and straightforward position determining modalities than that of hardware-based detection devices. The drawback of this approach is the bandwidth; however given the frequency response of nanoprobes as 30 Hz (Pollard et al., 2010), the 1 kHz achievable using this approach is more than adequate. Furthermore, more complex structures can be tolerated using an image-based approach, thereby offering further capabilities in the future. 4.2.4 Using FPGA Methods for Improved Bandwidth To increase the bandwidth of ROI detection, we also used a CMOS sensor with dedicated microelectronics for handling the very high image rate
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Fig. 6 User interface for camera control and analysis software. Centroid tracking is a simple procedure that can be implemented using control software such as LabView™. A 16 16 pixel region of interest can be used to achieve 1-kHz image acquisition and processing using an Orca 2.8 camera (Hamamatsu). The shape of the ROI has a little effect on the centroid capabilities, and therefore, this technique offers much greater flexibility in position detection.
(Towrie et al., 2009). The design of the CMOS camera chip allows parallel access to alternating pixel rows within microseconds. This permits up to 15k frames per second to be collected when taking ROI images. The workflow shown in Fig. 7 shows how a buffer created in dedicated electronics (known
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Field programmable gate array (operating frequency ~100 MHz) ROI Data Vanilla sensor output
Vanilla sensor control
Active pixel sensor ~20 k images/s
Data out (to PC)
ROI x,y coordinates FPGAPC link (Ethernet)
Sensor control signals
AOD movements
commands IN(from PC)
Data rate to PC ~20 MB/s
Output stage control
EOD
AOD
Acousto-optic deflector, repositioned at 60 kHz
Fig. 7 Data flow using an active pixel sensor approach to position detection. For much quicker position detection an active pixel sensor can be used. This device uses an FPGA to perform calculations on information as it leaves the sensor chip. This accelerates analysis because there is no requirement for data transfer to the PC for analysis there. As a result, this system can be used to feedback to the position of the traps for reduced noise if necessary. Image taken with permission from Towrie, M., Botchway, S. W., Clark, A., Freeman, E., Halsall, R., Parker, A. W., et al. (2009). Dynamic position and force measurement for multiple optically trapped particles using a high-speed active pixel sensor. The Review of Scientific Instruments, 80, 103704.
as a field-programmable gate array or FPGA) allows for robust communication between the PC and sensor, with image analysis performed on large image collections after the experiment. 4.2.4.1 Calibrating the Laser Tweezers
Two methods were used to measure trap stiffness, equipartition and Stokes’ drag force (Dupuis et al., 1997). Briefly, equipartition requires measuring the variance of a bead’s thermal motion while optically trapped using centroid detection: Trap stiffness ¼
kB T σ2
where kB is the Boltzmann constant, T the temperature of the fluid medium, and σ 2 the variance of the trapped bead (Dupuis et al., 1997). Measurement of trap stiffness using Stokes drag force provides independent confirmation of not only the stiffness but also the pixel calibration and centroid detection. For this an analog triangular wave is applied to the stage creating a constant velocity. This displaces the bead from the trap center
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resulting in a square wave output; the displacement from the center is used to calculate the restoring force of the trap because the viscous drag on the bead is easily calculated from the stage translation velocity: F ¼ 6πηrv where F is the force on the bead, η is the viscosity of the medium, r is the bead radius, and v is the constant stage velocity. Since the traps act like Hookean springs, their stiffness is easily calculated from: Trap stiffness ¼
F x
where x is the bead displacement from the trap center (Dupuis et al., 1997). In Fig. 8 we show results from both methods across a range of laser powers. This provides direct evidence that the detection system used is accurate. These methods have been well validated for spherical beads. The specific structure of the nanoprobe makes a highly precise measure of its
Fig. 8 Trap stiffness determined by equipartition and Stokes’ drag. Trap stiffness calculated using Stokes () and equipartition (□) is plotted against the applied laser power. A clear linear relationship is evident for both methods, and a strong similarity in stiffness across this range is used to validate this scheme of detection. Both calibrations were carried out on three separate beads. Standard deviation for each laser power is plotted as error bars (hard line for Stokes and dashed line for equipartition). Trap power is given at the source.
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hydrodynamic properties complex. However, it has been shown previously that when this structure is modeled as a large sphere its apparent trap stiffness is accurate enough to measure displacement forces on the nanoprobe (Pollard et al., 2010).
5. APPLYING THE NANOPROBE TO BIOLOGICAL STUDY SYSTEMS 5.1 Measuring the Tension of a DNA Tightrope With the nanoprobe detection and calibration established, it is now possible to study the properties of DNA tightropes. By deflecting the tightrope with the nanoprobe and measuring the position of the nanoprobe and the deflection of the tightropes, the pretension on the tightrope can be calculated. With the nanoprobe aligned tip toward the DNA and in the same z-plane, the stage is moved until contact with the DNA is observed. The stage is then programmed to move at a specific rate for a specific distance toward the DNA and then the position of the vertices measured (Fig. 9). The rate of movement can be slow; therefore, the CMOS detection approach is applicable. For the same stage movements, fluorescence images are also taken. These provide information on the amount of deflection of the tightrope and its initial length. The change in DNA length (Δl) is noted from the fluorescence image. The product of the trap stiffness and the displacement of the nanoprobe provides the total resistance to stretching offered by the DNA (Δf). We use the value for the increase in length at the peak force, since this is where the greatest displacement is seen. By simply calculating at what initial DNA extension a further increase in length Δl derives from a force of Δf using the worm-like chain extension model for DNA, it is possible to calculate the initial pretension on the DNA (Simons et al., 2015).
5.2 Measuring Interactions With Single Proteins One of the more sophisticated applications of this system is to study the interactions between proteins and DNA. This is achieved using a very similar setup to that described earlier. However, Qdot-labeled proteins are introduced prior to the addition of nanoprobes. The proteins will find their target sites on DNA, and these interactions can be probed physically. First, it is necessary to find a single DNA tightrope with a Qdot-labeled protein bound while carrying the nanoprobe in the optical traps. The nanoprobe
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Fig. 9 Using the nanoprobe to calculate DNA pretension. Fluorescence images of the nanoprobe are shown as it interacts with a DNA tightrope (A). The resulting bend in the DNA is used to calculate the change in contour length. The position of the nanoprobe is measured in X and Y for each vertex (B and C) and from this the force experienced as the nanoprobe is displaced from its trap center is calculated (D). Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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was found not to bind Qdots or Qdot-labeled protein (Simons et al., 2015). Therefore, its use in a protein-rich environment is not deleterious to its function as a probe, especially since the tip itself is not coated in protein. Once a Qdot-labeled protein is found attached to the DNA, it is possible to move the nanoprobe over the protein, as this occurs the nanoprobe will be held back by the protein until enough force is reached that it slips over the top (Fig. 10A). We performed similar tests with an immobile object and
Fig. 10 Force response of the nanoprobe to a protein bound to DNA. The DNA repair protein UvrA forms a dimer but for simplicity is termed UvrA rather than UvrA2; therefore, since we end-label the protein using a genetically encoded biotinylatable tag, we expect the attachment of two Qdots (Kad et al., 2010; Simons et al., 2015). This has been well characterized at the single-molecule level (Kad et al., 2010). Here we use the nanoprobe to understand how UvrA binds to DNA tightropes by measuring the resistance offered by the bound UvrA–Qdot conjugate to the passage of the nanoprobe. (A) Fluorescence images are shown of a nanoprobe interacting with a Qdot-labeled UvrA. (B) Plotting the peak forces as a histogram indicates two peaks which are interpreted to correspond to a topological map of the binding of UvrA–Qdot to the DNA tightropes. (C) Since the conjugate can bind in any orientation, the nanoprobe effectively probes all axial positions of the protein-Qdot by examining multiple molecules. Panel (A): Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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obtained very similar results (Simons et al., 2015). The force at which the nanoprobe slips over the top of the protein–Qdot conjugate is dependent on the size of the conjugate. By plotting a histogram of the peak force (Fig. 10B) a topological map of the protein–Qdot conjugate on the tightrope is obtained. In the case of UvrA, we see two peaks, likely corresponding to its asymmetric binding to DNA (Fig. 10C).
6. CONCLUSIONS AND OUTLOOK At present we are in a transformative era of the biosciences, where the complexity of systems is becoming apparent but at the same time tools from the physical sciences can offer new insights. Here we have presented an approach that uses a nanofabricated device to probe single protein molecules on single DNA tightropes. This has been used to map the binding of a protein to the DNA. Coupled with new probe designs that intelligently use the trapping field to imbue properties on the probe (Phillips et al., 2014), and high-speed imaging that records the probe movement and the corresponding single-molecule forces acting upon it with exquisite detail, the future holds promise for more elaborate and informative experiments. Furthermore, by conjugating the protein to the nanoprobe, it will be possible in the future to physically drag a protein along a tightrope, this means that the energy landscape of its binding can be discovered. For proteins that detect damage or specific sequences, we may be able to understand the energetics of these interactions. Such information is crucial if we want to be able to understand complex processes such as transcription initiation or DNA repair.
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Lyon, W. A., Fang, M. M., Haskins, W. E., & Nie, S. (1998). A dual-beam optical microscope for observation and cleavage of single DNA molecules and bond cleavage studies of single DNA molecules. Analytical Chemistry, 70, 1743–1748. Mehta, A. D., Finer, J. T., & Spudich, J. A. (1997). Detection of single-molecule interactions using correlated thermal diffusion. Proceedings of the National Academy of Sciences of the United States of America, 94, 7927–7931. Molloy, J. E. (1998). Optical chopsticks: Digital synthesis of multiple optical traps. Methods in Cell Biology, 55, 205–216. Molloy, J. E., Burns, J. E., Kendrick-Jones, J., Tregear, R. T., & White, D. C. (1995). Movement and force produced by a single myosin head. Nature, 378, 209–212. Noom, M. C., van den Broek, B., van Mameren, J., & Wuite, G. J. L. (2007). Visualizing single DNA-bound proteins using DNA as a scanning probe. Nature Methods, 4, 1031–1036. Otto, O., Gutsche, C., Kremer, F., & Keyser, U. F. (2008). Optical tweezers with 2.5 kHz bandwidth video detection for single-colloid electrophoresis. The Review of Scientific Instruments, 79, 023710. Phillips, D. B., Padgett, M. J., Hanna, S., Ho, Y.-L. D., Carberry, D. M., Miles, M. J., et al. (2014). Shape-induced force fields in optical trapping. Nature Photonics, 8, 400–405. Pollard, M. R., Botchway, S. W., Chichkov, B., Freeman, E., Halsall, R., Jenkins, D. W. K., et al. (2010). Optically trapped probes with nanometer-scale tips for femto-Newton force measurement. New Journal of Physics, 12, 113056. Qi, Z., Pugh, R. A., Spies, M., & Chemla, Y. R. (2013). Sequence-dependent base pair stepping dynamics in XPD helicase unwinding. eLife, 2, e00334. Riggs, A. D., Suzuki, H., & Bourgeois, S. (1970). Lac repressor-operator interaction. I. Equilibrium studies. Journal of Molecular Biology, 48, 67–83. Shon, M. J., & Cohen, A. E. (2016). Nano-mechanical measurements of protein-DNA interactions with a silicon nitride pulley. Nucleic Acids Research, 44, e7. Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486. Skinner, G. M., Baumann, C. G., Quinn, D. M., Molloy, J. E., & Hoggett, J. G. (2004). Promoter binding, initiation, and elongation by bacteriophage T7 RNA polymerase. A single-molecule view of the transcription cycle. The Journal of Biological Chemistry, 279, 3239–3244. Sung, J., Sivaramakrishnan, S., Dunn, A. R., & Spudich, J. A. (2010). Single-molecule dualbeam optical trap analysis of protein structure and function. Methods in Enzymology, 475, 321–375. Svoboda, K., Schmidt, C. F., Schnapp, B. J., & Block, S. M. (1993). Direct observation of kinesin stepping by optical trapping interferometry. Nature, 365, 721–727. Tafvizi, A., Huang, F., Leith, J. S., Fersht, A. R., Mirny, L. A., & van Oijen, A. M. (2008). Tumor suppressor p53 slides on DNA with low friction and high stability. Biophysical Journal, 95, L01–L03. Thompson, R. E., Larson, D. R., & Webb, W. W. (2002). Precise nanometer localization analysis for individual fluorescent probes. Biophysical Journal, 82, 2775–2783. Tokunaga, M., Imamoto, N., & Sakata-Sogawa, K. (2008). Highly inclined thin illumination enables clear single-molecule imaging in cells. Nature Methods, 5, 159–161. Towrie, M., Botchway, S. W., Clark, A., Freeman, E., Halsall, R., Parker, A. W., et al. (2009). Dynamic position and force measurement for multiple optically trapped particles using a high-speed active pixel sensor. The Review of Scientific Instruments, 80, 103704. van Loenhout, M. T., de Vlaminck, I., Flebus, B., den Blanken, J. F., Zweifel, L. P., Hooning, K. M., et al. (2013). Scanning a DNA molecule for bound proteins using hybrid magnetic and optical tweezers. PLoS One, 8, e65329.
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Integrating Optical Tweezers, DNA Tightropes, and Single-Molecule Fluorescence Imaging: Pitfalls and Traps J. Wang*, J.T. Barnett*, M.R. Pollard†, N.M. Kad*,1 *School of Biosciences, University of Kent, Canterbury, Kent, United Kingdom † DFM A/S, Kongens Lyngby, Denmark 1 Corresponding author: e-mail address: [email protected]
Contents 1. Introduction 2. Elongating Bundled DNA for Imaging 2.1 DNA Combing 2.2 DNA Tightropes 3. Integrating Laser Tweezers Into Biological Experiments 3.1 Introduction 3.2 Trapping With Nanoprobes 4. Controlling and Detecting the Nanoprobe 4.1 Using an AOD to Create Three Traps 4.2 Detection Strategies 5. Applying the Nanoprobe to Biological Study Systems 5.1 Measuring the Tension of a DNA Tightrope 5.2 Measuring Interactions With Single Proteins 6. Conclusions and Outlook References
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Abstract Fluorescence imaging is one of the cornerstone techniques for understanding how single molecules search for their targets on DNA. By tagging individual proteins, it is possible to track their position with high accuracy. However, to understand how proteins search for targets, it is necessary to elongate the DNA to avoid protein localization ambiguities. Such structures known as “DNA tightropes” are tremendously powerful for imaging target location; however, they lack information about how force and load affect protein behavior. The use of optically trapped microstructures offers the means to apply and measure force effects. Here we describe a system that we recently developed to enable individual proteins to be directly manipulated on DNA tightropes. Proteins bound to DNA can be conjugated with Qdot fluorophores for visualization and also Methods in Enzymology, Volume 582 ISSN 0076-6879 http://dx.doi.org/10.1016/bs.mie.2016.08.003
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directly manipulated by an optically trapped, manufactured microstructure. Together this offers a new approach to understanding the physical environment of molecules, and the combination with DNA tightropes presents opportunities to study complex biological phenomena.
1. INTRODUCTION Proteins and substrates exist in a physical milieu where they are subject to processes such as Brownian motion, diffusion, and external force. The contribution of these factors toward the overall mechanism requires the properties of the individual components to be determined for extrapolation to the ensemble system. One major biological process that requires understanding at this level is diffusion to target, and particularly so for targets on DNA. Based on diffusion models, it is possible to predict the association time of a protein to its target site (Berg, 1993). However, Lac repressor was found to locate its target site faster than predicted by a three-dimensional (3D) encounter model (Riggs, Suzuki, & Bourgeois, 1970). Facilitated diffusion was postulated to explain this discrepancy and involves a change in the mechanism of target search from being wholly 3D to also include onedimensional (1D) diffusion along the backbone of the DNA (Adam & Delbr€ uck, 1968; Winter, Berg, & von Hippel, 1981). For many years, numerous investigators set out to test this hypothesis (Halford & Marko, 2004) prior to the application of relatively new single-molecule methods. These methods permit the observer to track a single molecule labeled with a fluorophore as it moves toward its target site. Early investigations proved the existence of 1D sliding (Blainey, van Oijen, Banerjee, Verdine, & Xie, 2006; Tafvizi et al., 2008), hopping (Bonnet et al., 2008) which is longdistance jumps in position along a single DNA strand and also a combination of all of these mechanisms (Hughes et al., 2013; Kad, Wang, Kennedy, Warshaw, & van Houten, 2010). Also clear was that some proteins were capable of directional motor movement (van Oijen et al., 2003). To study such systems at the single-molecule level, first it requires DNA to be elongated. In Fig. 1 we show an image of a field of λ-DNA molecules stained with YOYO-1 dye. λ-DNA (the genome of bacteriophage lambda) consists of 48,502 base pairs; with a base pair separation of 0.34 nm, this equates to a contour length of 16.5 μm. It is apparent in Fig. 1 that the molecules in the image have collapsed into a bundled conformation. The radius pffiffiffiffiffiffiffiffi of a bundle can be calculated using random flight theory r ¼ l K ,
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Fig. 1 Image of YOYO-1 stained bacteriophage λ-DNA molecules. This DNA has a contour length of 16 μm; however, in solution, the DNA collapses into bundles. These diffuse through the field of view and are also seen to change shape.
where l is the contour length and K is the Kuhn length (twice the persistence length), for λ-DNA this is 1.3 μm. Comparing this to the point spread function of a typical fluorophore (Anderson, Georgiou, Morrison, Stevenson, & Cherry, 1992; Thompson, Larson, & Webb, 2002), it is apparent that measuring motion within this bundle is extremely challenging. Furthermore, the bundle will move, so the positional accuracy is adversely affected. Here we describe how we construct DNA tightropes to achieve elongated DNA structures and how this can be combined with optically trapped probes for more sophisticated experiments.
2. ELONGATING BUNDLED DNA FOR IMAGING 2.1 DNA Combing DNA can be laid onto an activated surface (Allemand, Bensimon, Jullien, Bensimon, & Croquette, 1997), usually polystyrene or polymethylacrylate by a method known as combing (Bensimon et al., 1994). This is achieved by either retracting an activated slide through a solution of DNA or flowing DNA across a surface (Crut et al., 2005; Deen et al., 2015; Lyon, Fang, Haskins, & Nie, 1998). Typically performed at low pH to achieve better binding, the DNA can then be examined using total internal reflection fluorescence microscopy. This reduces the amount of background fluorescence permitting fluorescently labeled proteins on DNA to be clearly imaged. In addition to this approach the DNA can be attached by one end and flow applied to elongate the DNA such as used to study the DNA repair protein
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human oxoguanine DNA glycosylase 1 (Blainey et al., 2006) or DNA replication (Lee et al., 2006). Although more complex, this approach reduces, but does not eliminate, interactions between DNA-bound proteins and the surface. Visually, however, it is unclear whether the proteins are attached to the DNA or surface because both are in the same focal plane.
2.2 DNA Tightropes This method exploits the simplicity of combing but raises the DNA from the surface without the need for flow. Suspending individual DNA molecules between surface-immobilized beads ensures that anything visible within the focal plane above the surface must be bound to DNA and not the surface. To facilitate imaging, it is necessary to optimize illumination such that only the tightropes are excited and the rest of the solution receives minimal light. This is achieved by using oblique angle fluorescence excitation microscopy (Hughes et al., 2013; Konopka & Bednarek, 2008; Tokunaga, Imamoto, & Sakata-Sogawa, 2008). To create DNA tightropes we use the following protocol: 1. Plasma clean 22 40 mm coverslips (Harrick PDC-32G, Ithaca, NY) for 2 min per slide. 2. Build a flow chamber as described in Fig. 2. 3. Block the chamber overnight by flowing in mPEG5000 (Sigma Aldrich) pH 8.2, wash with water, and then block overnight again with 50 mM Tris–HCl, 10 mg/mL BSA, 0.1% Tween20. 4. Flow in 5 μm silica beads (precoated overnight with 350 μg/mL poly-Llysine (Sigma Aldrich)), until bead separation is on average 20 μm. 5. Wash flow chamber with 200 μL buffer and check bead density again to ensure that beads have not been washed from the surface. In that event recoat beads with poly-L-lysine before adding to flow chamber (see Fig. 2 legend for details on constructing a flow chamber; also see Kad et al., 2010). 6. Attach a syringe pump (AL-1000 World precision instruments) to one flow chamber outlet and to the other a microcentrifuge tube using polyethylene tubing (PE90; BD Medical, Oxford, UK). 7. Backflow the buffer from the syringe into the microcentrifuge tube to ensure an airtight connection. 8. Flow in 100 μL of 15 nM λ-DNA, this will bring the DNA into the flow chamber. Apply forward and backward flow to run the DNA across the surface beads for at least 20 min.
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Fig. 2 Design of flow cells for creating tightropes. To create flow chambers a plasma cleaned glass coverslip (#1.5) is attached to a glass microscope slide using an in-house cut double-sided adhesive gasket (UK industrial tapes) of depth 180 μm. The glass slide is predrilled (using a diamond bur; Precision Dental, London, UK) to allow the attachment of small diameter tubing and sealed using UV curable adhesive (NOA68, Thorlabs, Ely, UK) after insertion. The gasket design is shown in the lower panel, indicating the positions of ports used for tightropes and for nanoprobe injection. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
9. Flush the flow chamber with 200 μL of imaging buffer and if required 100 μL of 1 nM YOYO-1 dye to visualize the DNA. During experiments, it is usual to exclude the addition of YOYO-1 dye since it is known that anything in focus at the height of the beads will be bound to DNA.
3. INTEGRATING LASER TWEEZERS INTO BIOLOGICAL EXPERIMENTS 3.1 Introduction Optical trapping, also known as laser tweezing, permits physical manipulation of single molecules. This approach requires a high-energy laser beam focused into a fluid sample to interact with particles of higher refractive index than their surrounding solution. As a consequence the particles, typically polystyrene or silica beads, become fixed near the focal point of the laser
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beam. Deviation from the focal point results in a restoring force that is proportional to the distance from the center of the trap position over a region of approximately 500 nm. This Hookean response results in a very simple linear correlation between displacement and force. Therefore, not only are laser tweezers useful for physical placement of molecules but also their response to force. The application of laser tweezers for studying molecular motors was initially established with kinesin (Svoboda, Schmidt, Schnapp, & Block, 1993) and myosin (Finer, Simmons, & Spudich, 1994; Guilford et al., 1997; Molloy, Burns, Kendrick-Jones, Tregear, & White, 1995). The latter experiments elegantly suspend individual actin filaments between optically trapped beads, pull them taut (Dupuis, Guilford, Wu, & Warshaw, 1997), and then use this platform to investigate the properties of single myosin molecules attached to a solid surface (Fig. 3). These experiments have revolutionized biology by permitting the load response of single molecules to be determined (Kad, Patlak, Fagnant, Trybus, & Warshaw, 2007; Kad, Trybus, & Warshaw, 2008; Veigel, Molloy, Schmitz, & Kendrick-Jones,
Fig. 3 The three-bead assay. This assay, used primarily for studies of the molecular motor myosin, involves plating a low density of myosin onto a coverslip surface that is precoated with 3-μm silica beads. 1-μm silica beads are trapped and used to capture the ends of an actin filament, which is then tautened. When the actin is brought into contact with the surface-bound myosin and the position of the trapped beads detected, a clear drop in variance is seen. This is due to the additional contribution of the stage to the stiffness of the system. In addition, myosin will move the actin which is seen as a deflection from the baseline of the bead position.
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2003; Veigel, Schmitz, Wang, & Sellers, 2005). Such measurements are not possible at the bulk level and also permit the contribution of the single molecule in the context of the ensemble to be determined. Such scaling measurements are integral to the construction of nanobiotechnological devices (Hess, 2011) and for the understanding of complex biological systems. Beyond molecular motors the study of DNA-based enzymes has also been facilitated by the use of laser tweezers. This has typically taken the form of suspending single DNA molecules between beads in much the same way as actin has been used for studying myosin (Hilario, Amitani, Baskin, & Kowalczykowski, 2009; Qi, Pugh, Spies, & Chemla, 2013; Skinner, Baumann, Quinn, Molloy, & Hoggett, 2004; Wang, Yin, Landick, Gelles, & Block, 1997). However, recently more elaborate applications of laser tweezers have been used to study the localization and affinity of proteins for DNA. These include looping one DNA molecule around another (Noom, van den Broek, van Mameren, & Wuite, 2007; van Loenhout et al., 2013) or incorporating atomic force microscope probes (Huisstede, Subramaniam, & Bennink, 2007; Shon & Cohen, 2016).
3.2 Trapping With Nanoprobes 3.2.1 Fabrication and Preparation of Nanoprobes Optical trapping has traditionally involved the use of spherical beads that are isotropically suspended in the traps. This means that the movement of the beads is limited to simple translation and that the surface available for interacting with molecules is large and cannot be easily controlled. To overcome these limitations, we and others have used structured particles that can be optically trapped. As shown in Fig. 4 we use a triangular structure with three
Fig. 4 The structure of the nanoprobe. Fabricated using EBL from SU8 the nanoprobe is a triangular structure with three cylinders located at each vertex. Three laser traps are used to position the probe each focused on a different cylinder. The probe is a long thin blade structure, and the nanoprobe is visible by fluorescence as well as bright-field microscopy. Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., Botchway, S. W., Parker, A. W., & Kad, N. M. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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trapping points and a protruding tip (Pollard et al., 2010). This permits accurate 3D positioning of the protrusion such that molecules can be manipulated with greater precision. The “nanoprobe” structures were fabricated using electron beam lithography as follows. To etch the nanoprobes a chrome layer followed by an SU8 photoresist layer is applied to a silicon wafer, baked, and then exposed (Pollard et al., 2010). This process yielded 4000 nanoprobes per segment on a silicon wafer. Correct release of the nanoprobes from the wafer is pivotal to enable subsequent experiments to be performed efficiently. First, the chrome layer is dissolved using ceric ammonium nitrate (CAN) pipetted directly onto the nanoprobe containing segment. The duration of exposure to etchant is controlled in order to leave some chrome connecting the nanoprobes to the wafer. This permits subsequent washing steps without loss of nanoprobes; any remaining CAN residue in the wafer region containing nanoprobes is gently washed away using high-purity water several times. Residual CAN was found to inhibit downstream steps, and washing also removes any free particulates that can contaminate the flow chamber and interfere with trapping experiments. After several washes the nanoprobes can be released with a more vigorous rinse with buffer containing BSA and Tween20 (Simons et al., 2015). This step prevents aggregation of the nanoprobes which are then collected and kept for use in the flow cell. 3.2.2 Flow Chamber Adaptations for Improved Experimental Yield For introduction of the nanoprobes into the flow chamber, it is necessary to avoid contact with the walls of the tubing, which was found to bind avidly to the nanoprobes. Therefore to add nanoprobes, a separate hole was drilled into the flow chamber requiring a specific design with a separate channel for nanoprobes (Fig. 2). This design not only avoids the use of polyethylene tubing during addition of nanoprobes but also creates a channel for storage of nanoprobes within the flow chamber. Nanoprobes are injected using a glass Hamilton syringe, and the channel can be sealed simply using commercially available sticky tape.
4. CONTROLLING AND DETECTING THE NANOPROBE Laser tweezers are relatively simple to construct, requiring only the back aperture of a high numerical aperture (>1.2) objective lens to be slightly overfilled with a stable laser. To obtain the correct beam size requires
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the use of one or more telescopes in the optical path. Controlling the position of the laser tweezers can be achieved by a number of schemes; however, they all share a common point that the turning mechanism is conjugated with the back focal point of the objective lens. This ensures a stable wide region for trapping, for a much more extensive discussion on the tweezer setup used (see Sung, Sivaramakrishnan, Dunn, & Spudich, 2010). Later we discuss the control mechanism used for our system (Simons et al., 2015) and the detection scheme.
4.1 Using an AOD to Create Three Traps In order to manipulate the laser-trapping beam laterally across the imaging plane, acoustic optical deflectors (DTD-274HD6M, IntraAction) are inserted into the laser beam path to deflect the beams with varying acoustic frequency (Molloy, 1998). In such a way, three optical traps can be created to coincide spatially with the vertex positions of the nanoprobe. These are generated by rapidly (40 kHz) repositioning the laser between each position significantly faster than the nanoprobe response frequency of 30 Hz (Pollard et al., 2010). Removal of the IR-blocking filter permits an image of the trap position to be projected onto the camera, although this should only be performed when necessary to prevent damage to the camera. Once a floating nanoprobe is found in solution, adjust the orientation of the traps to match it, and note their position. Turn off the IR-trapping laser. Move the trap position marker over the nanoprobe and adjust the focus to be slightly above the nanoprobe. Turning on the IR laser should capture the nanoprobe. Since the nanoprobes are located in the side channel of the flow chamber (Fig. 2), they will need to be navigated to an appropriate DNA tightrope. This is a delicate operation requiring movements over millimeters, and touching anything in the flow chamber will likely result in the nanoprobe sticking. At high IR laser powers, it is possible to guide the nanoprobe over the surface beads. However, to image the tightropes simultaneously, the trapping and imaging planes need to be offset in the z-axis. This is achieved by moving the focusing lens for the optical tweezers. Once a tightrope is found, the lens is returned to its correct position bringing the fluorescence and trapping planes into parfocality. At this point, it is necessary to align the detection apparatus with the center of the nanoprobe vertices and to perform some calibrations of the trap stiffness. For the latter, position detection is required.
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4.2 Detection Strategies Classical detection of trapped objects requires either a position sensitive detector (PSD) or a quadrant photodiode (QPD). These record the position of the trapped object at the back focal plane of the microscope condenser. The recorded signal is an interference signal between the scattered light from the bead and incident trapping beam. This detection approach relies on the linear relationship between the center of mass of the interference pattern and the relative position between the trapping beam and the trapped bead; however, this scheme is not suitable for tracking multiple beads simultaneously. The output of a PSD, usually a voltage signal, will be affected by parameters such as power of the beam and its shape and size; therefore, careful calibration is required. We adopt a more flexible detection method of bead tracking by using image-based position detection which allows us to track multiple beads by following their movement. Previous studies (Belloni, Monneret, Monduc, & Scordia, 2008; Keen, Leach, Gibson, & Padgett, 2007; Otto, Gutsche, Kremer, & Keyser, 2008) have shown that image-based tracking can provide similar performance compared with PSD and QPD detection schemes with the added advantage of multiple trap detection with flexible controls. Multiple moving objects can be tracked using real-time updated region of interests (ROIs) across the image. With PSD or QPD detectors offering up to hundreds of kHz, image-based trapping cannot compete in terms of detection bandwidth. However, for most position measurements, a fast camera with a carefully positioned ROI will suffice. 4.2.1 Optics for Image-Based Detection The position detection path and the laser-trapping beam are built around a wide-field microscope. As shown in Fig. 5, a high-power white LED light source is placed above the sample stage of the microscope in a transmission configuration to provide bright-field illumination. A Hamamatsu ORCA-Flash 2.8 CMOS camera (Hamamatsu Photonics, Hamamatsu, Japan) with 1920 1440 pixels is used as the image sensor to acquire brightfield microscopic images. The imaging signal collected by the microscope objective, OBJ, is focused by a doublet achromatic lens (L1, f ¼ 200 mm) to form a conjugate imaging plane where a mechanical pupil (A) is placed to control the aperture size. A pair of achromatic lenses L2 (f ¼ 100 mm) and L3 (f ¼ 50) form a 4F configuration to relay the image at A to the image sensor. The trapping laser is a 5W 1070-nm ytterbium fiber-coupled laser that is separated from the detection path by a dichroic beam splitter. The laser beam is focused by the objective to the sample plane, SP, within the
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Fig. 5 Layout of the camera-based position detection scheme. CMOS, Hamamatsu’s Scientific CMOS Camera, ORCA-Flash 2.8; DBS, dichroic beam splitter; A, aperture; L1, L2, and L3, achromatic lens with focal length of 200, 100, and 50 mm, respectively; LED, brightfield light source; NBF, notch-blocking filter; OBJ, microscope objective; SP, sample plane; the dashed arrows indicate the positions of image plane and image conjugate planes.
field of view of the camera. A notch-blocking filter is used to stop the back reflection of the trapping beam reaching the camera. In this way, the camera can deliver the image of the SP with the trapping laser focus in the middle. Each pixel on the camera covers an area of 120 120 nm2 in the SP; therefore, an ROI of 16 16 pixels produces a detection window of 1.92 1.92 μm2. With this ROI, the ORCA-Flash 2.8 camera runs at a maximum frame rate of 1 kHz. The pixel size is calibrated by using a graticule (Graticules Ltd., Tonbridge, UK). To initially locate objects for trapping the full field of view is used. ROIs are then adopted to isolate individual objects floating in the solution. 4.2.2 Centroid Detection We describe here the use of 1.5-μm diameter silica beads as a trapped object; however, the procedure for a nanoprobe vertex is identical. When the bead is trapped by the laser, the image of the bead appears bright in the middle, providing an image with high contrast and small size. The center of mass is calculated by using the centroid approach in LabView™. N X N X
Cx ¼
i I ij
i¼1 j¼1 N X N X I ij i¼1 j¼1
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The centroid position in x (Cx) takes the sum of the product of the intensity of each pixel in a column (Iij) and that column position in x (i). This is repeated for every column in the ROI (i ¼ 1 to N). Finally the sum of all of these values is expressed relative to total intensity of the image. This calculation is repeated for the y-axis but for each row. We normally threshold the image to offset problems of low contrast and to remove any distractions from background features in the flow chamber. Using this detection system clear steps (<15 nm, pixel size ¼ 120 nm) in either the “x”- or the “y”-direction of a 1.5-μm bead are visible when the sample is translated by a nanometer-accuracy piezo stage (LPS200 Mad City Labs). Statistical treatment of the data can provide much higher resolution, and when trapped, further correlation methods can be applied (Mehta, Finer, & Spudich, 1997). 4.2.3 Acquisition and Processing Software Two pieces of software were created to control the camera and analyze the data from an acquired image sequence, both written in LabView™ (National Instruments, Newbury, UK). The camera control software sets up the parameters for the CMOS camera, acquires, and can export the image data to a hard drive when the sequence is too large for system RAM (using 32-bit Windows XP PC this is limited to 3 GB). Centroid information for the trapped bead is concurrently reported in real time. A raw image of a 1.5μm bead is displayed Fig. 6 (top), and the centroid determined “x” and “y” positions are plotted alongside. Using off-line software (Fig. 6, bottom), it is possible to generate image intensity histogram information, apply new thresholds for recalculation of centroid coordinates values, and visualize the processed bead position with a two-dimensional mapping. This validated detection scheme offers the capability of using more versatile and straightforward position determining modalities than that of hardware-based detection devices. The drawback of this approach is the bandwidth; however given the frequency response of nanoprobes as 30 Hz (Pollard et al., 2010), the 1 kHz achievable using this approach is more than adequate. Furthermore, more complex structures can be tolerated using an image-based approach, thereby offering further capabilities in the future. 4.2.4 Using FPGA Methods for Improved Bandwidth To increase the bandwidth of ROI detection, we also used a CMOS sensor with dedicated microelectronics for handling the very high image rate
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Fig. 6 User interface for camera control and analysis software. Centroid tracking is a simple procedure that can be implemented using control software such as LabView™. A 16 16 pixel region of interest can be used to achieve 1-kHz image acquisition and processing using an Orca 2.8 camera (Hamamatsu). The shape of the ROI has a little effect on the centroid capabilities, and therefore, this technique offers much greater flexibility in position detection.
(Towrie et al., 2009). The design of the CMOS camera chip allows parallel access to alternating pixel rows within microseconds. This permits up to 15k frames per second to be collected when taking ROI images. The workflow shown in Fig. 7 shows how a buffer created in dedicated electronics (known
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Field programmable gate array (operating frequency ~100 MHz) ROI Data Vanilla sensor output
Vanilla sensor control
Active pixel sensor ~20 k images/s
Data out (to PC)
ROI x,y coordinates FPGAPC link (Ethernet)
Sensor control signals
AOD movements
commands IN(from PC)
Data rate to PC ~20 MB/s
Output stage control
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Acousto-optic deflector, repositioned at 60 kHz
Fig. 7 Data flow using an active pixel sensor approach to position detection. For much quicker position detection an active pixel sensor can be used. This device uses an FPGA to perform calculations on information as it leaves the sensor chip. This accelerates analysis because there is no requirement for data transfer to the PC for analysis there. As a result, this system can be used to feedback to the position of the traps for reduced noise if necessary. Image taken with permission from Towrie, M., Botchway, S. W., Clark, A., Freeman, E., Halsall, R., Parker, A. W., et al. (2009). Dynamic position and force measurement for multiple optically trapped particles using a high-speed active pixel sensor. The Review of Scientific Instruments, 80, 103704.
as a field-programmable gate array or FPGA) allows for robust communication between the PC and sensor, with image analysis performed on large image collections after the experiment. 4.2.4.1 Calibrating the Laser Tweezers
Two methods were used to measure trap stiffness, equipartition and Stokes’ drag force (Dupuis et al., 1997). Briefly, equipartition requires measuring the variance of a bead’s thermal motion while optically trapped using centroid detection: Trap stiffness ¼
kB T σ2
where kB is the Boltzmann constant, T the temperature of the fluid medium, and σ 2 the variance of the trapped bead (Dupuis et al., 1997). Measurement of trap stiffness using Stokes drag force provides independent confirmation of not only the stiffness but also the pixel calibration and centroid detection. For this an analog triangular wave is applied to the stage creating a constant velocity. This displaces the bead from the trap center
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resulting in a square wave output; the displacement from the center is used to calculate the restoring force of the trap because the viscous drag on the bead is easily calculated from the stage translation velocity: F ¼ 6πηrv where F is the force on the bead, η is the viscosity of the medium, r is the bead radius, and v is the constant stage velocity. Since the traps act like Hookean springs, their stiffness is easily calculated from: Trap stiffness ¼
F x
where x is the bead displacement from the trap center (Dupuis et al., 1997). In Fig. 8 we show results from both methods across a range of laser powers. This provides direct evidence that the detection system used is accurate. These methods have been well validated for spherical beads. The specific structure of the nanoprobe makes a highly precise measure of its
Fig. 8 Trap stiffness determined by equipartition and Stokes’ drag. Trap stiffness calculated using Stokes () and equipartition (□) is plotted against the applied laser power. A clear linear relationship is evident for both methods, and a strong similarity in stiffness across this range is used to validate this scheme of detection. Both calibrations were carried out on three separate beads. Standard deviation for each laser power is plotted as error bars (hard line for Stokes and dashed line for equipartition). Trap power is given at the source.
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hydrodynamic properties complex. However, it has been shown previously that when this structure is modeled as a large sphere its apparent trap stiffness is accurate enough to measure displacement forces on the nanoprobe (Pollard et al., 2010).
5. APPLYING THE NANOPROBE TO BIOLOGICAL STUDY SYSTEMS 5.1 Measuring the Tension of a DNA Tightrope With the nanoprobe detection and calibration established, it is now possible to study the properties of DNA tightropes. By deflecting the tightrope with the nanoprobe and measuring the position of the nanoprobe and the deflection of the tightropes, the pretension on the tightrope can be calculated. With the nanoprobe aligned tip toward the DNA and in the same z-plane, the stage is moved until contact with the DNA is observed. The stage is then programmed to move at a specific rate for a specific distance toward the DNA and then the position of the vertices measured (Fig. 9). The rate of movement can be slow; therefore, the CMOS detection approach is applicable. For the same stage movements, fluorescence images are also taken. These provide information on the amount of deflection of the tightrope and its initial length. The change in DNA length (Δl) is noted from the fluorescence image. The product of the trap stiffness and the displacement of the nanoprobe provides the total resistance to stretching offered by the DNA (Δf). We use the value for the increase in length at the peak force, since this is where the greatest displacement is seen. By simply calculating at what initial DNA extension a further increase in length Δl derives from a force of Δf using the worm-like chain extension model for DNA, it is possible to calculate the initial pretension on the DNA (Simons et al., 2015).
5.2 Measuring Interactions With Single Proteins One of the more sophisticated applications of this system is to study the interactions between proteins and DNA. This is achieved using a very similar setup to that described earlier. However, Qdot-labeled proteins are introduced prior to the addition of nanoprobes. The proteins will find their target sites on DNA, and these interactions can be probed physically. First, it is necessary to find a single DNA tightrope with a Qdot-labeled protein bound while carrying the nanoprobe in the optical traps. The nanoprobe
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Fig. 9 Using the nanoprobe to calculate DNA pretension. Fluorescence images of the nanoprobe are shown as it interacts with a DNA tightrope (A). The resulting bend in the DNA is used to calculate the change in contour length. The position of the nanoprobe is measured in X and Y for each vertex (B and C) and from this the force experienced as the nanoprobe is displaced from its trap center is calculated (D). Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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was found not to bind Qdots or Qdot-labeled protein (Simons et al., 2015). Therefore, its use in a protein-rich environment is not deleterious to its function as a probe, especially since the tip itself is not coated in protein. Once a Qdot-labeled protein is found attached to the DNA, it is possible to move the nanoprobe over the protein, as this occurs the nanoprobe will be held back by the protein until enough force is reached that it slips over the top (Fig. 10A). We performed similar tests with an immobile object and
Fig. 10 Force response of the nanoprobe to a protein bound to DNA. The DNA repair protein UvrA forms a dimer but for simplicity is termed UvrA rather than UvrA2; therefore, since we end-label the protein using a genetically encoded biotinylatable tag, we expect the attachment of two Qdots (Kad et al., 2010; Simons et al., 2015). This has been well characterized at the single-molecule level (Kad et al., 2010). Here we use the nanoprobe to understand how UvrA binds to DNA tightropes by measuring the resistance offered by the bound UvrA–Qdot conjugate to the passage of the nanoprobe. (A) Fluorescence images are shown of a nanoprobe interacting with a Qdot-labeled UvrA. (B) Plotting the peak forces as a histogram indicates two peaks which are interpreted to correspond to a topological map of the binding of UvrA–Qdot to the DNA tightropes. (C) Since the conjugate can bind in any orientation, the nanoprobe effectively probes all axial positions of the protein-Qdot by examining multiple molecules. Panel (A): Image taken with permission from Simons, M., Pollard, M. R., Hughes, C. D., Ward, A. D., van Houten, B., Towrie, M., et al. (2015). Directly interrogating single quantum dot labelled UvrA2 molecules on DNA tightropes using an optically trapped nanoprobe. Scientific Reports, 5, 18486.
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obtained very similar results (Simons et al., 2015). The force at which the nanoprobe slips over the top of the protein–Qdot conjugate is dependent on the size of the conjugate. By plotting a histogram of the peak force (Fig. 10B) a topological map of the protein–Qdot conjugate on the tightrope is obtained. In the case of UvrA, we see two peaks, likely corresponding to its asymmetric binding to DNA (Fig. 10C).
6. CONCLUSIONS AND OUTLOOK At present we are in a transformative era of the biosciences, where the complexity of systems is becoming apparent but at the same time tools from the physical sciences can offer new insights. Here we have presented an approach that uses a nanofabricated device to probe single protein molecules on single DNA tightropes. This has been used to map the binding of a protein to the DNA. Coupled with new probe designs that intelligently use the trapping field to imbue properties on the probe (Phillips et al., 2014), and high-speed imaging that records the probe movement and the corresponding single-molecule forces acting upon it with exquisite detail, the future holds promise for more elaborate and informative experiments. Furthermore, by conjugating the protein to the nanoprobe, it will be possible in the future to physically drag a protein along a tightrope, this means that the energy landscape of its binding can be discovered. For proteins that detect damage or specific sequences, we may be able to understand the energetics of these interactions. Such information is crucial if we want to be able to understand complex processes such as transcription initiation or DNA repair.
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