Open-frame system for single-molecule microscopyAdriel Arsenault, Jason S. Leith, Gil Henkin, Christopher M. J. McFaul, Matthew Tarling, Richard Talbot,Daniel Berard, Francois Michaud, Shane Scott, and Sabrina R. Leslie Citation: Review of Scientific Instruments 86, 033701 (2015); doi: 10.1063/1.4913271 View online: http://dx.doi.org/10.1063/1.4913271 View Table of Contents: http://scitation.aip.org/content/aip/journal/rsi/86/3?ver=pdfcov Published by the AIP Publishing Articles you may be interested in A bisected pupil for studying single-molecule orientational dynamics and its application to three-dimensionalsuper-resolution microscopy Appl. Phys. Lett. 104, 193701 (2014); 10.1063/1.4876440 Combined versatile high-resolution optical tweezers and single-molecule fluorescence microscopy Rev. Sci. Instrum. 83, 093708 (2012); 10.1063/1.4752190 Multispot point spread function for multiphoton fluorescence microscopy Rev. Sci. Instrum. 80, 096104 (2009); 10.1063/1.3226658 Diffusion of carbon nanotubes with single-molecule fluorescence microscopy J. Appl. Phys. 96, 6772 (2004); 10.1063/1.1815053 Visualization of individual carbon nanotubes with fluorescence microscopy using conventional fluorophores Appl. Phys. Lett. 83, 1219 (2003); 10.1063/1.1599042

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REVIEW OF SCIENTIFIC INSTRUMENTS 86, 033701 (2015)

Open-frame system for single-molecule microscopyAdriel Arsenault, Jason S. Leith, Gil Henkin, Christopher M. J. McFaul, Matthew Tarling,Richard Talbot, Daniel Berard, Francois Michaud, Shane Scott, and Sabrina R. Lesliea)

Department of Physics, McGill University, Montreal, Quebec H3A 2T8, Canada

(Received 22 September 2014; accepted 10 February 2015; published online 2 March 2015)

We present the design and construction of a versatile, open frame inverted microscope systemfor wide-field fluorescence and single molecule imaging. The microscope chassis and modulardesign allow for customization, expansion, and experimental flexibility. We present two componentswhich are included with the microscope which extend its basic capabilities and together create apowerful microscopy system: A Convex Lens-induced Confinement device provides the system withsingle-molecule imaging capabilities, and a two-color imaging system provides the option of imagingmultiple molecular species simultaneously. The flexibility of the open-framed chassis combinedwith accessible single-molecule, multi-species imaging technology supports a wide range of newmeasurements in the health, nanotechnology, and materials science research sectors. C 2015 AIPPublishing LLC. [http://dx.doi.org/10.1063/1.4913271]

I. INTRODUCTION

Since its introduction, fluorescence microscopy hasserved as a work horse for molecular and cell biology. A widerange of synthetic fluorescent dyes have been developed, andmany fluorescent proteins that are expressed directly withinhost cells have been isolated and modified.1 The ample numberof options means that nearly any biomolecular target can belabelled and observed using a microscope.

New single-molecule microscopy techniques haveemerged in line with the development of fluorophores, in-cluding confocal,2,3 total internal reflection (TIRF),4 and two-photon microscopy.5 These approaches allow informationnormally hidden within statistical ensembles—such as an indi-vidual molecule’s structure, position, or state—to be probeddirectly. Single-molecule experiments have led to criticaldiscoveries spanning the fields of physics,6,7 chemistry,8,9

biology,10–13 and materials science.14,15 The creation of super-resolution techniques has further extended the resolution offluorescence microscopy beyond the diffraction limit.16 Thesuccess of these techniques has ensured that they will remainvaluable tools for researchers and continue to furnish newdiscoveries.

While there is no shortage of commercially availablemicroscopes to choose from, there are significant limitations tousing most “off-the-shelf” microscope and imaging systems.Many have a closed-box design that fundamentally limits theways in which they can be used. This often makes it difficultto develop custom experimental devices that integrate with thesystem. To overcome this challenge, we present a versatile,open-frame, inverted fluorescence microscope system, whichincludes a laser excitation system, a dual-emission imagingsystem, and a Convex Lens-induced Confinement (CLiC)device. The open frame allows for the introduction of newdevices, facilitates diagnostics, and allows further modular

a)sabrina.leslie@mcgill.ca. URL: http://www.physics.mcgill.ca/leslielab/.

additions to be made independently. For any researcher wish-ing to implement any or all of the following infrastructure fortheir own purposes, schematics, CAD files, and a full list ofall commercial parts will be made available by contacting thecorresponding author.

II. CUSTOM MICROSCOPE

Our inverted microscope chassis is shown in Fig. 1. Itsopen-frame concept allows for integration with experimentaldevices and customization of optical components. The micro-scope is the base unit for the microscopy system presented inthis work. While it was designed to integrate with the CLiC andtwo-colour imaging system, it also functions independently asan inverted fluorescence microscope.

The microscope chassis consists of three plates con-structed from aluminum tooling plate for precisely parallelfaces. These bottom and middle plates are separated by four1.5 in.-diameter stainless steel pillars, and the middle and topare separated by four 1 in.-diameter stainless steel pillars.The bottom plate serves as a base for the structure. The topof the center plate supports the dichroic cube holder assembly.The dichroic holder functions by allowing an adjustable sliderto lock in place, which is compatible with most commerciallyavailable mounted dichroics. A mirror and tube-lens assemblyattaches to a 90◦ adjustable turret that is mounted on the bottomof the center plate. The rotating turret allows the user to directthe fluorescence to one of two imaging systems: either directlyto an EMCCD camera or to a two-color imaging system(outlined in Sec. IV). The top plate supports the objectivefocusing assembly as well as an X-Y translation stage. Theobjective is mounted on a motorized lens positioner, whichallows for precise focusing. For a list of all custom machinedparts see Table I.

The modular design of this microscope allows for vari-ability in features and cost. For example, if the option to switchemission pathways quickly is not required, the rotating turret

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FIG. 1. 3D schematic of the microscope chassis. The labelled parts are asfollows: (i) objective; (ii) X-Y stage; (iii) objective collar; (iv) motorized lenspositioner; (v) dichroic; (vi) rotating mirror; (vii) tube (with tube lens); (viii)dichroic slider mount. For a full list of custom parts see Table I.

can be removed. Additionally, an optical encoder used for pre-cise measurements of the objective’s position is recommendedfor many applications but not required for basic microscopy.

III. EXCITATION OPTICS

A simplified schematic of the excitation pathway is shownin Fig. 2. The beams exit the three lasers (488 nm, 561 nm,and 647 nm) before two long-pass dichroics combine them intoa single beam. Once combined, the beam passes through twosets of telescopes that each consists of two lenses separated bytheir back focal lengths. Together, the two telescopes expandthe beam by a factor of ∼45. The first telescope is made fromlenses with focal lengths of 50 mm and 150 mm and the secondfrom lenses with focal lengths of 50 mm and 750 mm. Thismagnification allows the lasers to uniformly illuminate the fullfield of view.

Finally, the combined beams pass through the wide-fieldlens, which focuses the light on the back focal plane of theobjective. The wide-field lens is mounted on a micrometer-driven stage that serves as the base for the final mirror andlens assembly. This assembly controls the position at whichthe beam hits the dichroic and, ultimately, the angle at whichit falls incident on the sample, allowing TIRF microscopy tobe performed.

This excitation setup allows the use of high-poweredlasers (∼120 mW) to excite single molecules allowing forshort exposure times. The beam pathway was simulated and

TABLE I. Custom components of chassis.

Description Quantity Materials

Aluminum base plate 3 Aluminum tooling plateRotation turret 1 Brass, 6061 aluminumDichroic holder assembly 1 6061 aluminumTube lens support 1 6061 aluminum

optimized using OSLO optical design software. Optics werechosen to minimize the chromatic focal shift between the threelasers. The final setup has a maximum focal shift of 0.01 mmbetween the blue and green lasers and 0.56 mm between theblue and red lasers.

The open-frame concept of the excitation system allowsfor significant customization. For example, the lasers can beblocked independently using shutters placed in front of thecombining dichroics or shared between two separate micro-scopes through the use of a beam splitter. More components,such as an acousto-optic tunable filter (AOTF), can be addedinto the excitation pathway to allow for techniques such asFluorescence Resonance Energy Transfer with AlternatingLaser Excitation (FRET-ALEX),17 or super resolution tech-niques such as Stochastic Optical Reconstruction Micros-copy (STORM).16 The customizability of the microscope alsomakes it suitable for use with multiple imaging techniques atonce (e.g., combining fluorescence with polarization opticsto incorporate measurements of molecular orientations androtational dynamics18).

IV. DUAL-CHANNEL IMAGING SYSTEM

For many experiments, it is crucial to be able to simul-taneously label and image more than one molecular speciesusing spectrally distinct probes. As is the case with mostcommercially available microscope chassis, two-color imag-ing systems often have a closed-box design which typicallyrestricts the choice of spectra and may not allow for the spectraof each channel to be controlled independently during anexperiment. The custom dual-emission imaging system thatwe have created as part of our microscopy system overcomesthese limitations through its open and customizable design.

Light collected from the objective can be sent to the dual-emission imaging system by rotating the lens tube using itsrotation turret. This setup allows for quick and easy switchingbetween single-view and dual-view experiments. Our dual-emission imaging station (Fig. 2, 1-12) is made from simple,off-the-shelf components. The system uses optical elementswith a 2 in.-diameter in order to reduce aberrations and customemission filters to control the spectra viewed in each channel.

The dual-emission system functions by creating two spec-trally distinct images of the same field of view and allowsthem to be viewed simultaneously on the same CCD chip.The dual imaging system achieves this by placing a physicalslit at the first imaging plane. This slit cuts the image to halfthe size of the camera’s CCD chip (8.12 mm by 4.06 mm foran Andor iXon camera). Wavelengths above and below thedichroic’s characteristic wavelength are directed to separateoptical paths, forming two spectrally distinct images. Thelong-wavelength image is passed through one path, while theshort-wavelength image is reflected down the other. The choiceof the dichroic’s transmission spectrum depends on the flu-orophores being imaged. The transmitted and reflected pathseach contain two identical lenses which are separated by theirfocal lengths. The system is designed to use two lenses to focusthe image rather than one, as our simulations showed that thissetup reduces chromatic and spherical aberrations. Each path

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FIG. 2. Excitation and dual-emission optical pathways: The top pathway is a simplified schematic of the excitation optics. a-j represent the emission pathwaywhere the laser beams are combined and expanded before entering the microscope. 1-12 represent the dual-emission imaging pathways where the image is splitaccording to wavelength and recombined side-by-side on the camera. For a complete parts list of both pathways see Table II. Note: the two systems are notshown to scale with respect to each other.

has a single mirror that allows the two images to be positionedside-by-side before they are recombined into a single beamby a second dichroic. For a list of all components used in theexcitation and emission pathways, see Table II.

The dual-emission imaging system can be easily modifiedto allow the use of a wide range of fluorophores by plac-ing the dichroics on magnetic mounts, allowing them to beeasily exchanged between experiments. The open design ofthe dual imaging system also allows for emission filters tobe placed in specific imaging pathways rather than havingmany two-band pass emission filters in a conventional dichroicturret. The emission filters can also be mounted on sliders,allowing for quick insertion or removal of the filters (Fig. 2).This is particularly useful for experiments performed usingCLiC microscopy where interferometry (direct imaging ofthe exciting laser) is used to measure the chamber’s heightprofile.20–23

TABLE II. List of optical components shown in Figure 2.

Excitation pathway Emission pathways

Position Description Position Description

a 561 nm dichroic 1 Adjustable mechanical slitb 488 nm dichroic 2 640 nm dichroicc Filter wheel 3 Lensd 1st lens of 1st telescope 4 Emission filter (on slider)e 2nd lens of 1st telescope 5 Lensf 1st lens of 2nd telescope 6 2 in. mirrorg 2nd lens of 2nd telescope 7 2 in. mirrorh Widefield lens 8 Lensi Custom microscope 9 Lensj EMCCD camera 10 Emission filter

11 640 nm dichroic12 EMCCD camera

V. SINGLE-MOLECULE IMAGING WITH CLIC

Single-molecule microscopy has enabled researchers tovisualize the dynamics, conformational states, and multi-species interactions characterizing a wide range of systems,yielding mechanistic insights which were previously inacces-sible using ensemble-averaged measurement techniques.6–15

In this section, we outline a modular device which extends thecapabilities of our microscope to single-molecule microscopyusing CLiC.20–23

CLiC imaging is based on a simple working principle:confining molecules to a thin sample chamber to allow forsingle-molecule imaging. CLiC microscopy is performedwhen a curved optical lens (referred to as the “push-lens”)pushes into and deforms the top coverslip of a flow cell. Thecoverslip bows into the sample chamber creating a thin wedge-shaped chamber that constrains molecules within a single focalplane. The confined volume improves the signal-to-noise ratioas the image is not contaminated with light emitted by out-of-focus molecules from above or below. Molecules confined inthe chamber are free to diffuse within the focal plane and can betypically tracked for tens of seconds within a 100 × 100 µm2

field of view. When compared with other single-moleculemicroscopy approaches such as confocal and TIRF, CLiCoffers extended observation times and allows a greater rangeof molecular concentrations.20–23

Figure 3 demonstrates the flow cell CLiC device presentedin this work. This device represents a simplified version of aCLiC device which is presented in Ref. 23. Unlike the pre-vious device which operates through piezo actuators and hasan intricate fluidics delivery system, this device is manuallycontrolled which means it is both easy to construct and simpleto use, and has the further additional benefit of being finan-cially accessible. The simple design also allows for easier cus-tomization of experiments. While this device lacks the extremeprecision of the computer controlled device, we demonstrate in

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FIG. 3. 3D model of the manual flow cell CLiC device. (a) Top view of the device. The four largest holes are for fastening the plate onto the microscope. Theclamps can be positioned on either set of holes in the central part of the plate depending on the flow cell size. (b) Cross-section of the CLiC device. The levercan be displaced by turning the lock-nut, which rotates on a 1/4-100 in.-threaded screw. (c) Close-up of the cross-section at the contact point between the flowcell and the push-lens. The displayed top and bottom coverslips are, respectively, 145 µm and 175 µm thick. The oil objective is typically positioned 170 µmaway from the bottom of the bottom coverslip. Reprinted with permission from C. M. J. McFaul et al., “Single-molecule microscopy using tunable nanoscaleconfinement,” Proc. SPIE 8811, 881102 (2013). Copyright (2013) by Leslie Lab and SPIE.

Sec. VI B that we are able to take data which is of comparablequality to previous devices.

The device rests atop a sample plate that is bolted to themicroscope stage. Custom aluminum fittings hold a steel shaftthat acts as a rotation-axis for an extended lever arm. On thislever, the push-lens is fixed in a recess by means of set screws.The lever is initially raised to allow for sample insertion andlowered by adjusting a nut on a finely threaded rod at theopposite end of the lever. As the nut is tightened towards theplate, the lens presses down on the flow cell, which is held overthe imaging aperture by custom spring clips. A spring applies arestoring force that ensures gradual chamber compression andremoves the backlash in the adjustment nut.

VI. PERFORMANCE OF SYSTEM AND APPLICATIONS

A. Single-molecule photobleaching

In order to demonstrate a number of the versatile capa-bilities of our microscopy system, we have taken a seriesof demonstration data. To demonstrate the single-moleculeimaging capabilities of the microscope, we observed strep-tavidin singly labelled with Alexa Fluor 488 fluorescent dye(Life Technologies). The molecules were diluted to 180 pM inphosphate buffered saline solution (PBS), with a pH of 7.74.The streptavidin molecules were bound to the bottom coverslipof the flow cell using a polyethylene glycol (PEG) coatingcomprising 1% biotinylated PEG.24

A flow cell was formed with 30 µm-thick double-sidedtape with the PEG-coated coverslip on the bottom, and a cover-slip cleaned with piranha solution (3:1 H2SO4: 30% H2O2)on the top. While being observed on the microscope, labelledstreptavidin was flowed into the sample chamber and allowedto bind with the biotin. After 10-15 min, excess streptavidinwas washed out. While the bottom coverslip was in focus,the laser beam angle was altered until total internal reflectionoccurred and single molecules could be seen clearly. Video

was taken at 200 ms exposure and 7.5 mW of 488 nm laserpower. A cropped image (115 × 115 pixels) of streptavidinmolecules taken through the short path of the dual-emissionimaging system can be seen in Fig. 4(a).

Observing intensity traces of individual particles showedclear photobleaching steps. Figure 4(b) shows the intensityof the single molecule within the red circle shown in (a) vs.time, normalized by the background intensity. The moleculewas observed for 400 frames at 200 ms/frame. The moleculebleaches at frame ∼160 after which the intensity drops to thelevels found at background, which verifies that the moleculeis singly labelled. The point spread function (PSF) of thismolecule was analyzed in (c) and (d): the former is the rawintensity data of the individual molecule, and the latter is thisdata fit to a Gaussian function. The Gaussian fit has a standarddeviation of 0.62 ± 0.8 pixels in X and 0.58 ± 0.8 pixels in Y.For particles emitting at 525 nm, and a 60X objective with anumerical aperture (NA) of 1.49 (Nikon Apochromat TIRF),the Abbe diffraction limited radius is ∼0.66 pixels.

B. Particle tracking

In order to demonstrate the unique imaging capabilitiesthe CLiC device grants our microscopy system, we trackedsingle freely diffusing λ-phage DNA molecules. We comparedthe diffusion coefficients of these molecules at two differentregions of confinement within the sample chamber, demon-strating CLiC as a platform for nanoscale confinement spec-troscopy.

Figures 5(a) and 5(b) present fluorescent images offreely diffusing λ-phage DNA molecules stained with YOYO-1 fluorescent dye. The flow cell chamber heights for themolecules shown and analyzed lie between 460–640 nmand 1100–1470 nm. For a discussion on how these chamberheights are determined using a CLiC device see Ref. 23. Thelarger height is approximately equal to the bulk radius ofgyration, Rg , of the molecules. Polymer chains generally,25

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FIG. 4. (a) Image of a 115×115 pixel field of view of streptavidin molecules fluorescently labelled with Alexa Fluor 488 bound to a biotinylated-PEG coatingof a coverslip. (b) A background normalized intensity vs. time plot of a singly-labelled molecule (circled in red in (a)) which undergoes a single photobleachingstep. (c) Raw intensity data of the molecule highlighted in (a). (d) Raw data shown in (c) fit to a Gaussian function with standard deviation of 0.6 and 0.5 pixelsin x and y, respectively.

FIG. 5. Diffusion coefficients and in-plane radius of gyration of λ-phage DNA molecules. (a) and (b) Images of fluorescently labelled DNA molecules at heightsof H≈ 550 nm and H≈ 1280 nm, respectively. (c) and (d) Histograms of diffusion coefficients, D, for freely-diffusing molecules. The mean values (designatedby red lines) are 0.18±0.02 µm2/s (550 nm) and 0.23±0.02 µm2/s (1280 nm). (e) and (f) Histograms of in-plane radius of gyration, R∥. Mean values are1.20±0.22 µm (550 nm) and 1.1±0.14 µm (1280 nm). Reprinted with permission from C. M. J. McFaul et al., “Single-molecule microscopy using tunablenanoscale confinement,” Proc. SPIE 8811, 881102 (2013). Copyright (2013) by Leslie Lab and SPIE.

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and DNA in particular,26 have been shown to experiencethe effects of confinement at heights . 2Rg . Therefore, themolecules are confined for both ranges of chamber heightswith stronger confinement effects at the smaller chamberheight. The corresponding trajectory analysis and the distribu-tion of diffusion coefficients, D, shown in Figs. 5(c) and 5(d),demonstrate the effect of the imposed confinement in slowingDNA diffusion. Similarly, at these heights, we measured thein-plane radius of gyration:

R∥ =

i(ri − r̄)2Ii

i Ii, (1)

which measures the square root of the mean squared distancefrom the centroid of the particle, r̄, weighted by the pixels’intensities, I. Figures 5(e) and 5(f) demonstrate an increasein the in-plane radius of gyration with greater confinement.All molecules were tracked for a total of 55 s (1000 frames).Figures 5(c) and 5(d) are based on 66 and 92 lifetime-weightedparticles, respectively. For an overview of the analysis proto-col, we used to identifying and track particles, see Ref. 23.

For the preparation of the sample shown in Fig. 5, thefollowing experimental methods were followed. The final con-centrations of reagents were 1.445 mM Tris base, 0.445 mMboric acid, 0.1 mM EDTA, 0.32 mM HCl, 577 nM AlexaFluor 647, 285 mM 2-mercaptoethanol, and 13 pM ofYOYO-1 fluorescently stained λ-DNA at a labeling ratio ofone fluorophore per 10 base pairs. This solution had a finalpH of 7.1 and an ionic concentration of 1.35 mM. Cover-slips for all experiments were cleaned using 2:1 sulfuric acidto 30% hydrogen peroxide solution (piranha) for 45 min.The coverslips were then treated with 1M KOH for 15 min,rinsed thoroughly with deionized water, and dried before beingassembled into a flow cell.

C. FRET with dual-channel imaging

To demonstrate the capabilities of the dual-channel imag-ing system, we present a basic Förster Resonance EnergyTransfer (FRET) experiment. FRET is a technique which usesa pair of complementary fluorophores to detect when twomolecules have reached close physical proximity (∼10 nm orless).27 This technique works for fluorophore pairs in whichthe “donor” fluorophore’s emission spectrum overlaps withthe absorption spectrum of the “acceptor” fluorophore. Forsuitable fluorophores, energy from the donor fluorophore istransferred to the acceptor fluorophore through dipole-dipoleinteractions. This allows the acceptor to fluoresce even thoughit does not receive energy directly from the excitation laser.

In our demonstration experiment, small oligonucleotidesare bound to a chamber that is passivated with PEG, as inSec. VI A. Short oligonucleotides were bound to this sur-face through a biotin-streptavidin bond. These DNA segmentswere labeled with Cy5 (depicted in Fig. 6(a)). Once bound,complementary oligonucleotides which were labeled with Cy3were flowed into the flow cell. These complementary oligosselectively bound to the immobilized oligos in such a way asto bring the Cy3-Cy5 FRET pair into close proximity. Thesample was excited with green laser light (561 nm) and a FRET

FIG. 6. FRET Assay using bound oligonucleotides labelled with Cy5 andcomplementary oligo labelled with Cy3. (a) represents oligos bound to thePEGylated surface with a streptavidin-biotin bond and excited with a 561 nmlaser. (b) represents the above after the complementary oligos have beenintroduced. (b)-i represents two oligos which bind and produce a FRET sig-nal, while (b)-ii represents an oligo with no compliment and only fluorescesas in (a). In (c) a 140 by 100 region is shown where three bound moleculesare fluorescing in the green channel, and one in the red channel. The yellowcircles correspond to single oligos shown in (b)-ii, while the green circle cor-responds to a FRET pair as in (b)-i. (d) shows the mean counts of the particlecircled in green over 550 frames in both the green and red channels. In bothchannels the intensity drops to background in a single frame demonstratingthat the FRET donor fluorophore has bleached.

signal was observed in a fraction of the molecules. A schematicof this FRET interaction between bound oligos is shown inFig. 6(b).

The dual-channel imaging system allows for viewing boththe donor and acceptor fluorophores simultaneously in sepa-rate channels. Figure 6(c) shows a small region (140 by 100pixels) of both channels. Within this region immobile oligosthat do not have a complementary oligo bound to them fluo-resce only in the “green” channel (the molecules circled inyellow in Fig. 6(c)) and schematically depicted in 6(b)-ii, whilethose that do have a bound oligo fluoresce in both channels(circled in blue and depicted in 6(b)-i).

Molecules which showed a FRET signal had their inten-sities tracked over time and were found to photobleach inboth channels at the same frame. These instances were takento represent single donor fluorophore photobleaching, which

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simultaneously extinguishes the acceptor. The lifetime inten-sities of the two fluorophores in Fig. 6(c) showing this phenom-enon (circled in blue) are shown in Fig. 6(d). Signal cross-talkbetween channels was shown to be insignificant as the oligoswithout FRET pairs (yellow circles) did not appear in the redchannel.

The exact procedure for the experiment was as follows:Streptavidin in a buffer of 10 mM Tris-HCl, 100 mMNaCl, and 0.05% Tween-20 was flowed into the cham-ber, allowed to sit for 5 min, and then washed out with200 µl of the buffer. The Cy5 labelled oligo (5′-/5Cy5/ACCTCGCGACCGTCGCCA/3BiodT/-3′, purchased fromIDT), at a concentration of 10 pM in a conjugation buffer of10 mM Tris-HCl, 1.0 nM EDTA, and 2.0M NaCl was flowedin, allowed to incubate, and then washed out with 200 µlof 10 mM Tris-HCl. Lastly, the complementary oligo witha Cy3 label, (5′-TGGCGACGGTCGCGAGGT/3Cy3Sp/-3′,also purchased from IDT) was flowed in at a concentrationof 1 nM in 10 mM Tris-HCl. The reagents were allowed toincubate for 15 min, and then washed out with 400 µl ofbuffer. 10 mM TrisHCl including protocatechuic acid andprotocatechuate-3,4-dioxygenase, a deoxygenation system,was flowed in and allowed to act on the buffer in darkness for30 min. Images were acquired using total internal reflectionillumination with 300 EM gain and a 100 ms exposure time.

VII. CONCLUSION

We have presented the design and construction of a versa-tile open-frame fluorescence microscopy system and havedemonstrated a number of applications for wide-field andsingle-molecule fluorescence experiments. We have shown oursystem’s dual-channel imaging system allows for imaging be-tween spectrally distinct channels, and that our manual CLiCdevice is a powerful tool for single-molecule and confinementexperiments of freely diffusing molecules. Together thesefeatures can combine for novel experiments.

The microscopy system’s open frame offers many advan-tages to systems which are sold as sealed “black boxes.” Thedesign of the microscope and optical pathways allows accessto the beam at all points, easing troubleshooting and setup, andallows for the customization and easy expansion of the system.

ACKNOWLEDGMENTS

The authors would like to thank the agencies who havecontributed funding for this project: the Natural Sciences andEngineering Research Council of Canada NSERC, the Can-ada Foundation for Innovation (CFI), and the Department ofPhysics, McGill University. Adriel Arsenault, Shane Scott,and Dr. Jason Leith would like to thank the Cellular Dy-namics of Macromolecular Complexes Program (CDMC) fortheir graduate and postdoctoral fellowships. Daniel Berard andGil Henkin are also grateful for their graduate fellowships

awarded from the NSERC CREATE training program in Bio-nanomachines. Chris McFaul would like to thank both Bio-nanomachines and CDMC, and Daniel Berard would like tothank Bionanomachines and the McGill Science Undergrad-uate Research Awards (SURA) for summer research fellow-ships which helped them to make contributions to this work.

We are also grateful to the µManager software and devel-opment team as well as Andrew Caleb Guthrie, AlexanderVerge, and Yang Zhou for their help to develop our partic-ular implementation of the software. We would also like tothank Pascal Bourseguin and Steve Kacani for their role inmachining our custom pieces. We would finally like to thankSPIE for permission to reproduce selected material which waspresented in a conference proceeding.19

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