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Ultramicroscopy 110 (2010) 844–851

Contents lists available at ScienceDirect

Ultramicroscopy

0304-39

doi:10.1

n Corr

E-m

journal homepage: www.elsevier.com/locate/ultramic

Molecular recognition of DNA–protein complexes: A straightforward methodcombining scanning force and fluorescence microscopy

Humberto Sanchez a,n, Roland Kanaar a,b, Claire Wyman a,b

a Department of Cell Biology and Genetics, Cancer Genomics Center, Erasmus MC, PO Box 2040, 3000 CA Rotterdam, The Netherlandsb Department of Radiation Oncology, Erasmus MC, PO Box 2040, 3000 CA Rotterdam, The Netherlands

a r t i c l e i n f o

Article history:

Received 22 September 2009

Received in revised form

11 January 2010

Accepted 16 March 2010

Keywords:

Scanning force microscopy

Fluorescence microscopy

DNA–protein complex

Single-molecule studies

Fluorescent probes

91/$ - see front matter & 2010 Elsevier B.V. A

016/j.ultramic.2010.03.002

esponding author. Tel.: +31 10 704 3158; fax

ail address: h.sanchezgonzalez@erasmusmc.n

a b s t r a c t

Combining scanning force and fluorescent microscopy allows simultaneous identification of labeled

biomolecules and analysis of their nanometer level architectural arrangement. Fluorescent polystyrene

nano-spheres were used as reliable objects for alignment of optical and topographic images. This

allowed the precise localization of different fluorescence particles within complex molecular assemblies

whose structure was mapped in nanometer detail topography. Our experiments reveal the versatility of

this method for analysis of proteins and protein–DNA complexes.

& 2010 Elsevier B.V. All rights reserved.

1. Introduction

Imaging processes at the single-molecule level reveals infor-mation otherwise inaccessible by ‘‘bulk’’ experiments whereindividual features are usually hidden by ensemble averaging.Studying individual molecules allows observation of intermedi-ate–transient states in dynamic reactions, extraction of preciseinformation from asynchronous mixtures and determination ofthe proportion of different heterogeneous forms within a popula-tion. Scanning force microscopy (SFM, also known as atomic forcemicroscopy or AFM) is one such single molecule tool thatproduces images of biomolecules with nanometer resolution inthe absence of external contrast agents. Three-dimensionalinformation is collected from ‘‘soft’’ biological matter like proteincomplexes or nucleic acids likely without affecting relevantconformation and retaining overall native structure (for exampleas shown in Ref. [1]). In the last two decades, a wealth ofinformation about different proteins and their interactions withDNA has been produced using this technique [2–7]. Proteins caneasily be distinguished from DNA. Their influence on DNAstructure can be quantitatively determined from such imagesdue to the distinct shape and the well described mechanicalproperties of DNA [8]. However in complex assemblies it is notoften possible to distinguish proteins from each other based onshape and size. Several advances in SFM techniques are aimed atovercoming this limitation. Because the interaction force varies

ll rights reserved.

: +31 10 704 4743.

l (H. Sanchez).

between different materials, the oscillations of the cantilevercould be used to identify different molecules. For instance usingintermittent contact mode imaging the phase signal can be usedto produce surface hydrophobicity maps [9]. Another recentlydeveloped approach can identify different materials based onresonance frequencies of the cantilevers as in ‘‘bimodal atomicforce microscopy’’ [10]. Molecular recognition with functionalizedtips has been reported for the identification of specific proteins(for a review see Ref. [11]) but involves a trade off with decreasedspatial resolution as coated tips become larger and less welldefined. Although, the above-mentioned methods have partiallysolved the molecular recognition problem, they all lack one of themost attractive features of conventional SFM: its simplicity.

In biological imaging, specific molecules are often identified byattaching different florescent dyes or objects. Fluorescencedetection methods thus allow molecular identification based onemission wavelength. Current technical advances in opticalimaging allow detection down to the level of single fluorophoresand can report on multiple fluorescent properties. Time resolutionin the order of nanoseconds allows the study of dynamic processof individual molecules. However, due to Rayleigh criterion,spatial resolution is limited in the best case to half of theexcitation wavelength. Super-resolution beyond the diffractionlimit, down to nanometer precision, can be achieved bytechniques such as stimulated emission depletion (STED) micro-scopy [12], photoactivated localization microscopy (PALM) [13]and stochastic optical reconstruction microscopy (STORM) [14].Super-resolution of many fluorescent dyes is possible withSTED although the best resolution requires depletion intensitiesas high as GW/cm2. PALM and STORM require photo-switchable

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H. Sanchez et al. / Ultramicroscopy 110 (2010) 844–851 845

fluorophores but only need intensities in the range of W/cm2.These advanced optical methods can localize fluorophores withnm precision; however even at hypothetical atomic resolu-tion, structural details would still be limited to the labeledcomponents.

A combination of SFM and fluorescence microscopy mergesimaging capabilities to overcome limitations of the individualtechniques. This also has the advantage of relatively simple SFMand standard optical imaging. Several reports have alreadydescribed the development and use of such combined platforms.Identification of fluorescence markers on cell membranes ormeasuring adhesion forces constitute the main applications[15–17]. Other dedicated set-ups have probed their utility incombining fluorescence spectroscopy and SFM for identifyingDNA and polystyrene beads [18], lipid organization [19] or lightharvesting complexes in membranes [20]. However, imaging ofbiomolecules such as DNA–protein complexes remains techni-cally challenging because the objects of interest are smaller thantypical optical resolution. One of the main challenges of combin-ing SFM and fluorescence imaging is the unambiguous assign-ment of a fluorescence signal (typically a few hundrednanometers wide) to a globular structure of a few nanometers,like a protein complex. Although triangulation based on thecenter of fluorescence of quantum dots can be useful [21],blinking, photobleaching or heterogeneous excitation of isolatedfluorophores make the correlation with topography difficult.

We show here nanometer localization of single particles usinga combined scanning force and fluorescence microscope. The setup is based on a conventional wide-field fluorescence microscopeusing a mercury arc lamp combined with common excitation andemission filters. Intermittent contact mode scanning force micro-scopy in air was done with a commercially available scannercoupled to an optical microscope. Our experiments reveal theversatility of this method for recognition and simultaneouslocalization of different fluorescent-tagged objects, proteins andprotein–DNA complexes. We describe the use of fluorescencepolystyrene nano-spheres as reliable objects for alignment ofoptical and topographic images. This allowed us to preciselylocalize different fluorescent particles with nanometer resolutionwithin complex molecular assemblies, whose structure wasmapped in nanometer detail topography.

2. Materials and methods

2.1. Reagents

We used the following fluorescent dyes and objects: Fluo-Spheress carboxylate-modified microspheres (0.04 mm diameter,yellow-green fluorescent (505/515), orange fluorescent (540/560)and red fluorescent (580/605) from Invitrogen), semiconductornanocrystals streptavidin coupled Qdot525 and Qdot655 (‘‘quan-tum dots’’) (Invitrogen) and Alexa-488 (495/519) dye, AlexaFluor-633 (632/647) conjugated streptavidin (Invitrogen).

2.2. Sample preparation

Linear dsDNA (2054-bp) with a single 50 biotin was generatedby PCR amplification of pBluescript DNA using a modifiedbiotinylated oligo essentially as described previously [22].Nucleoprotein filaments, RAD51 coated dsDNA, were made byincubation of l DNA (Roche) and Alexa-488 conjugated humanRAD51 as previously described [23].

Unless otherwise stated, the indicated amount of product wasdiluted in 20 mL of deposition buffer (10 mM HEPES-KCl, pH 7.5,

10 mM MgCl2). Samples were deposited over freshly cleaved mica(muscovite V-1 quality, from Electron Microscopy Science)mounted on glass cover slips (24 mm round from Menzel-glazer)with optical adhesive NOA88 (Norland products) cured with UVlight (350 nm) for 3 min at room temperature. Before gluing, themica disc was cleaved with tape until almost transparent. Then aminimum amount of optical adhesive was put on the cover slip.The mica disc was attached and, by pressing down on it, the lowviscosity adhesive was homogeneously spread. After curing theadhesive and before using the mica-cover slip, the mica surfacewas made as thin as possible, by cleaving as many layers aspossible off with tape. This produces a mica glued glass substratethat allows focusing on the sample, with the objective lens wehave used here that has a 170 mm working distance the mica layerwill be 30 mm or less. After 1 min the sample drop was rinsedwith MilliQ water and dried with filtered air. Before depositingthe sample, autofluorescence on the mica surface was eliminatedby reduction with a drop of sodium borohydride [24] (0.25% w/v)for 20 min at room temperature and rinsed with MilliQ water.

2.3. Fluorescence microscopy

Samples were observed with a Nikon TE 2000U microscopyusing a Nikon 60� (NA 1.45) TIRF oil objective and collected by aCascade II:512B EMCCD camera (Princeton Instruments) drivenwith MetaMorph software (Molecular Devices). Excitation of thefluorophores was performed with a mercury arc lamp and stan-dard filter cubes (center wavelength/bandwidth, excitation-emis-sion, in nm): FITC (480/30-535/40) and TxRed (560/40-630/60)from Nikon, and GFP-3035B (472/30-520/35) from Semrock.

2.4. Scanning force microscopy

Scanning force microscopy was done with a NanoWizardsIIscanner (JPK instruments) mounted on the fluorescence micro-scope. Air dried samples were scanned in intermittent contactmode (air). Silicon Tapping/Non-Contact Mode tips 125 mm inlength with a spring constant of 25–75 N/m were from AppliedNanostructures. Drive frequency of the cantilevers used was onaverage 300 kHz. Images were, in general, acquired at 2 Hz linerate. Correlation of fluorescence and topographic images wasaccomplished first by the JPK DirectOverlayTM software thenrefined to nanometer accuracy as describe in results making useof structures with distinct identifiable shape and fluorescencesuch as nano-spheres.

3. Results

3.1. Identification and nanometer localization of individual

fluorescent particles

In order to validate precision alignment of optical andtopographic images to nm accuracy, we used commerciallyavailable fluorescently labeled polystyrene-beads, referred to asfluorospheres. With these we could distinguish three differentfluorescence particles with a combined SFM/fluorescence micro-scope. Such objects are needed as ‘‘fiduciary markers’’ or referencepoints in these combined images. Yellow-green (505/515), orange(540/560) and red fluorescent (580/605) 40 nm diameter beadswere mixed (60 pM each) and sonicated in a water bath for30 min. A solution (3 pM each) was diluted and deposited on aglued-mica glass surface as described in the experimental section.Samples were first observed with the inverted epifluorescencemicroscope. Fluorescent emission signals from ‘‘green’’ and ‘‘red’’

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beads were recorded after filtering the emission spectrum withFITC (480/30-535/40) and TxRed (560/40-630/60) cubes, respec-tively. A high numerical aperture objective (1.45) with a shortworking distance (0.17 mm at 23 1C) was used. So, mica thicknesswas limited to 30 mm excluding the glass. We did not observe areduction of fluorescence intensity due to the presence of themica compared to glass in these conditions. Nor were birefrin-gence effects observed (Supplementary Fig. 1). Lenses and mirrorsguiding the light produce a variable degree of optical aberrationsreflected in the final image. The DirectOverlayTM software allowsthe automatic calibration of the optical images by mapping theactual position of the cantilever (determined by the piezoprecision and not by the optical pathway) at 25 differentpositions. We calibrated regions of 40�40 mm, that are extra-polated to cover 50% of the 60�60 mm surface, in eachexperiment. This allowed us to precisely correlate SFM imageswith optical images of 512�512 pixels (�100 nm/pixel using a60� objective). However, because the initial position of thecantilever is unknown, there is an offset of a few microns in X andY between the optical and topographic images, as shown inFig. 1A. When imaging structures at the single molecule level(nm), it becomes apparent that in the absence of a clear reference

Fig. 1. Unique identification of different fluorophores by combined optical and

scanning force microscopy: (A) optical image of a mixture of three different

fluorospheres: red, green and orange. Green and red channels were overlaid.

Orange polystyrene beads are recognized by the colocalization of both signals. The

indicated area (green frame) was chosen for SFM; (B) alignment procedure using

fluorospheres. Red fluorescent signal emitted from the polystyrene beads shows

an offset in X and Y coordinates after the image calibration procedure; (C) height

features in topographic image (scale as in panel E) are easily aligned with the

fluorescent pattern. Overlay of the red channel with the SFM image is shown;

(D) green channel and the overlay with the SFM image and (E) height image.

(For interpretation of the references to coluor in this figure legend, the reader is

referred to the web version of this article.)

the offset will result in a serious disorientation and inability toidentify specific objects. We used the pattern of randomlydeposited fluorospheres evident in both topographic and opticalimages for alignment as shown in Fig. 1. The red fluorescencepattern had an X, Y offset (�4.8, �3.6 mm) with respect to theheight image (Fig. 1B). After alignment of the red spot mass-centers, with the height structure mass-centers both picturescorrelate with nanometer accuracy (Fig. 1C). Overlapping red andgreen signals identified the orange beads as shown in Fig. 1A.The boxed area in Fig. 1A was selected for SFM scanning, shownin Fig. 1E. The separate red and green emission images areoverlaid on the topography (Fig. 1C and D) for correlationand identification of two red, two green and one orange sphere.Volume analyses of the particles (calculated as previouslydescribed [6]) indicated the presence of one or more sphere perfluorescence spot in this example (Fig. 2B). Intensity measure-ments from non-saturated images where plotted against their

Fig. 2. Linear correlation between fluorescence intensity and particle volume:

(A) Volume in arbitrary units (a.u.) [6] versus integrated fluorescence intensity

(also in a.u.) [23] coming from the same regions were plotted and analyzed by

regression fit. The red and green lines and squares represent data from individual

particles red fluorospheres and green fluorospheres, respectively. Note that some

particles are smaller than complete fluorospheres and some represent more than

one sphere, in the range of particle sizes shown here and (B) fluorescence intensity

distribution of the spheres. Intensity bin size equivalent to fluorescence from one

fluorosphere (volume 1.971.2�105 a.u., N¼22), representing 3.5�105 a.u.

(green) and 8�105 a.u. (red). Nred¼95, Ngreen¼99. (For interpretation of the

references to coluor in this figure legend, the reader is referred to the web version

of this article.)

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respective volume revealing a linear correlation (Fig. 2A).Different fluorescence objects were identified in a complexmixture based on emission wavelength and their intensitycorrelated with size or number of fluorescent particles.

3.2. Topography reveals a large fraction of dark quantum dots

We similarly characterized quantum dots by depositing amixture of green Qdots and red fluorospheres or red Qdots andgreen fluorespheres (semiconductor nanocrystals streptavidin

Fig. 3. Fluorospheres as molecular fiduciary markers: (A) optical image showing red flu

region in A and height images overlaid; (C) red channel picture from the same region an

fluorospheres and red quantum dots: height image and red fluorescence channel overla

from panel (E); height image and fluorescence channels overlaid showing the double na

fluorescence. Total pixel intensity in regions around the indicated two red dots in pane

time of 30 s. Scale bar 2 mm. (For interpretation of the references to coluor in this figu

coupled Qdot525 and Qdot655, Invitrogen). To account forquantum dot blinking, consecutive images were collected andcompiled into a single average composite (500 ms expositiontime, 60 frames, 30 s total exposure time per image). Fig. 3Ashows the final Qdot image overlaid with signal from thefluorospheres. The indicated areas were selected for topographicimaging (Fig. 3B and C). Fluorescence signal from the fluoro-spheres could be overlaid onto structures with the expecteddimensions (�40 nm height). There was a homogenouspopulation of smaller structures corresponding to the size of

orospheres and green quantum dots; (B) green channel picture from the selected

d height images overlaid; (D) enlarged area from white rectangle in (B); (E) green

id. Qdots analyzed in panel (G) are labeled by numbers 1 and 2; (F) zoomed region

ture of one specific red fluorescent spot (labeled 2) and (G) time traces of Qdot655

l (E) (1 and 2) were measured from 60 frames acquired for 500 ms each in a total

re legend, the reader is referred to the web version of this article.)

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Fig. 4. SFM topographic image of DNA at nanometer resolution and localization of attached fluorescent label: (A) optical image showing the fluorescence signal from red

fluorospheres; (B) green quantum dots associated with biotinylated DNA, overlaid with the height image; (C) zoomed overlay region marked in B and (D) height image, Z

scale 0–2 nm, dark to bright. Scale bar 0.5 mm. (For interpretation of the references to coluor in this figure legend, the reader is referred to the web version of this article.)

H. Sanchez et al. / Ultramicroscopy 110 (2010) 844–851848

Qdots (between 5 and 10 nm height). Around 10% (17 fluorescentobjects out of 160 countable structures in the best case likeFig. 3B) of these Qdot sized entities were detected in theircorresponding fluorescence channel (green in Fig. 3A, B and D;and red in Fig. 3E and F). The presence of a majority of Qdot sizedstructures that were not fluorescent is similar to a previous report[21]. A detailed look at the height images revealed additionalinformation difficult to obtain from fluorescence alone. Forinstance, intensity over time traces for the two red Qdots (1 and2) highlighted in panel 3E shows blinking behavior as expected(Fig. 3G). Although there was no clear signature that allowed us toascribe this fluorescence behavior to more than one fluorophore, azoomed scan of the selected area (Fig. 3F) revealed the presence oftwo Qdot sized structures very close together contributing to thisone fluorescent spot.

3.3. Fluorescent labeled DNA molecules

We produced nm resolution images of DNA identified by afluorescent label coupled specifically to one end. For identificationand localization of specifically labeled proteins in complexassemblies with DNA it will be impossible to correlate topographyand fluorescence for structures smaller than optical resolutionwithout a marker object. DNA can be fluorescently labeled and isoften made visible by intercalating dyes [25,26]. However, suchdyes distort DNA structure and frequently interfere with protein–DNA binding activity. Here, in order to identify DNA by afluorescent marker we use a fluorescent label coupled specificallyto the end of DNA molecules.

Streptavidin coupled quantum dots (Qdot525) were attachedto linear 2054-bp DNA molecules that included one 50-biotin. ThisDNA was deposited for imaging together with red fluorospheres(3 pM). Green and red signals, from the Qdots and the fluoro-spheres, respectively, were recorded separately (Fig. 4). Theoverlay of topography again indicated most Qdots were notfluorescent in the conditions used (Fig. 4C). However, DNAmolecules can be clearly identified in topography by length

(�0.6 mm) and width (�10 nm) most with Qdots at one terminus(Fig. 4D). Note that more than one DNA could attach to a Qdotbecause streptavidin is multivalent (glowing or not).

3.4. Detection of fluorescent proteins

To demonstrate nm resolution imaging of individual proteins wedeposited fluorescent streptavidin (red) together with fluorospheres(green) and first detected emission signals from both fluorophores(Fig. 5A). A region including both streptavidin and fluorospheresignals was selected for topography imaging. The 60-kD streptavidinprotein is expected to be approximately a 5 nm diameter sphere,smaller than the 40 nm polystyrene fluorospheres. In addition, eachstreptavidin (reported to have 4 dye molecules per tetramer,molecular probes-Certificate of analysis, or our measurement of2.7) will have far fewer dye molecules than the fluorospheres(3.5�102 fluorescein equivalents in the yellow-green fluorospheres,molecular probes). In the zoomed images (Fig. 5B, C and D) thefluorospheres were distinctly identifiable by both fluorescence andtopography. There were in addition many small objects visible intopography that corresponded to the size of streptavidin. Smallerobjects were proven not to be scanning noise by shifting the SFMscan direction and observing the same constellation of objects (datanot shown). With the simple optical imaging we applied here, mostof the single proteins did not show obvious fluorescence abovebackground. After using the fluorospheres to align the optical andtopography images, the red fluorescence signal could be identifiedcorresponding to a complex likely including several streptavidinproteins, larger than the predominant single protein molecules. Thuswe are able to distinguish individual protein molecules and observefluorescence for complexes corresponding to several proteins.

3.5. Visualizing DNA–protein complexes

Our eventual aim is to identify proteins in complex structuresbased on different fluorescent labels. As an example of this type ofanalysis we used a version of the human RAD51 protein labeled

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Fig. 6. Fluorescence human RAD51 protein filaments on DNA: (A) optical image showing green fluorescence filamentous structures originated by the association of the

Alexa 488 conjugated RAD51 with the long DNA molecules of lambda phage (�48 kbp). Height image overlaid with the optical image from red fluorospheres; (B) or green

RAD51; (C) from the same region shown in A; (D) overlay of height and green fluorescence images of one filament end and (E) extra resolution in the height image shows a

lariat at the end of this filamentous structure. Scale bar 2 mm. (For interpretation of the references to coluor in this figure legend, the reader is referred to the web version of

this article.)

Fig. 5. Localization of individual fluorescence streptavidin particles: (A) optical image showing the fluorescence signal from green fluorospheres and AlexaFluor-633

conjugated streptavidin (in red); (B) and (C) Selected area in A shows individual fluorescence channels overlaid with the height image and (D) height image alone. Scale bar

0.5 mm. (For interpretation of the references to coluor in this figure legend, the reader is referred to the web version of this article.)

H. Sanchez et al. / Ultramicroscopy 110 (2010) 844–851 849

with a single fluorophore in a defined position [23]. AlexaFluor488 labeled RAD51 filaments covering double-strandedDNA, 48 kbp phage lambda genome, were mixed with redfluorospheres and deposited for imaging. Green and red channelswere individually recorded and overlaid after acquisition (Fig. 6).Green filamentous structures are DNA covered with RAD51. Inthis case we could detect the presence of the protein by itscolor and additionally define the structure of the filamentsby the topography. The topography revealed structural detail,indistinguishable in the optical image, such as multiple filamentsclose together (Fig. 6B and C) and the filament end curled into aloop (Fig. 6D and E). Here the DNA filament is obvious in bothoptical and topographic images due to the fluorescent protein.However, it is clear that even non-fluorescent proteins and bareDNA would be visible for analysis in such topographic images,where the relative location of labeled proteins within complexassemblies could be accurately determined.

4. Discussion

We describe here the use of a commercial instrument andsimple sample preparation methods for simultaneous optical andtopographic imaging of nano-objects, proteins and DNA. Biomo-lecules such as DNA, proteins and their complexes were depositedon mica as they are for many standard SFM imaging experiments.The methods described avoid chemical cross-linking of thecomplexes to be observed thus avoiding possible structuralartifacts this can cause. In addition we use untreated surfaces topreserve DNA conformation and allow quantitative measurementof changes induced by proteins [8]. In contrast, a recent similardemonstration of combined SFM/optical imaging [25] relied ontreated surfaces and DNA stretching methods that would precludeanalysis of important protein-induced changes in DNA conforma-tion that are often of interest. Topographic images of biomole-cules such as DNA and proteins are also more reliable on

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atomically flat mica surfaces compared with much rougherglass.

We present here several improvements over the previouslyreported combined SFM-fluoresence imaging of protein–DNAcomplexes that make our set-up more efficient. We showlabeled-RAD51 protein associated with unlabeled DNA (Fig. 6).The fluorescence and height images are correlated using fluoro-spheres added for this purpose. Moreover, using the same imagewe show the position of fluorospheres of different color andtheir correlated height image (Figs. 1 and 2). We addressed thesuitability of Qdots for correlating topography and fluorescenceby studying the dark fraction and observe a high dark fractionmaking them unacceptable for this purpose (Fig. 3). DNA labeledat one end by Qdots was imaged by SFM showing the expectedheight, width, shape and resolution for DNA. The fluorescent end-bound Qdots could be identified. We show a small protein that ishard to unambiguously identify in topography, Alexa fluor 633labeled-streptavidin but can be identified in clusters by fluores-cence (Fig. 5). In all of the presented cases, the correlation oftopography and fluorescence relies on the use of fluorospheresand not on the fortuitous emergence of discernable shape in thesample objects themselves, and is therefore generally applicableto aligning similar images for any biomolecules. Anotherimportant advantage of our set up is the use of 25 pictures ofthe cantilever that are used by the manufacturer’s software tocalibrate the optical image and determine the position of the tipin the topographical images, avoiding in this way, opticalaberrations. We are currently using this approach to analyzecomplexes involving several proteins acting together on DNA inthe DNA repair processes we study. Identification of the relativeposition of specific proteins will provide a wealth of informationabout their functional arrangements.

Many applications that we envision combining optical andSFM topographic imaging will involve objects that are smallerthan the resolution limit of standard optical imaging. In order toaccurately map optical signals and topography, both the availablecalibration of optical images by the JPK DirectOverlayTM systemand fine tuning offset adjustment to align optical and topographicimages to nm precision are needed to obtain useful information.We demonstrate here the use of fluorescent polystyrene nano-spheres as convenient fiduciary objects for this purpose. Thefluorospheres attach to mica together with the biomoleculesduring standard deposition and fall at random on the surface. Anyimage including at least 3 fluorospheres can be accuratelycorrected for offset by aligning the nanometer resolutiontopography images of the spheres with the center of the opticalspots. Thus fluoropsheres are robust, convenient, economical andnon-interfering markers for coordinated SFM and optical imaging.We used this alignment to identify clusters of fluorescentstreptavidin among the multitude of monomeric streptavidindetected in the topographic image. Correlating topography andoptical images also revealed that under the conditions we use amajority of quantum dots are not fluorescent, and thus cannot beused as robust fiduciary markers. Furthermore, it was possible toaddress unambiguously the proportion of dark particles in oursemiconductor nanocrystals. Fluorescence correlation spectro-scopy (FCS) studies [27] reported a quantum dot preparation as44% dark, 36% blinking and the remaining fraction was assumedto be glowing. In our experimental set up individual particlescould be observed indefinitely with an equivalent dwell time of500 ms. We did not observe a glowing fraction but a non-darkfraction that was entirely blinking. However, likely due to limitedfreedom of the particles in ambient humidity conditions theproportion of the dark fraction was higher than expected fromsolution. Fluorescence properties of quantum dots are verysensitive to surface changes [28]. Our data support the idea that

the quantum yield reported for the particles is proportional to thenon-dark fraction. Recently, the generation of non-blinking Qdotsthat would overcome this limitation has been reported [29]. Otherfluorophores and defined molecular complexes such as Alexa dyecoupled proteins with known stoichiometry will have to beanalyzed to determine if the surface and imaging conditions ingeneral influence the efficiency of detecting fluorescent mole-cules. For instance the Alexa labeled RAD51, already characterizedto include one fluorophore per monomer [23,30], may be usefulfor characterizing our sample preparation and imaging systemwith respect to this feature.

With a relatively simple optical set up and samples preparedon a mica–glue–glass substrate we have not yet demonstratedsingle fluorophore sensitivity. However this sensitivity is onlylimited by the sophistication of the optical set up and analysis.The number of different fluorophores that can be detected islimited by the filters installed and can easily be increased, forinstance by exciting the quantum dots with shorter wavelengthsfrom the arc lamp spectrum or by using a dedicated laser. Inaddition, higher power laser excitation of specific fluorophoreswill increase detection sensitivity. Similarly, nanometer localiza-tion of single fluorophores should also be possible with suffi-ciently bright or accumulated emission signals. Using a highermagnification objective (100� ) and the calibration of a smallerarea of 30�30 mm will increase the localization accuracy of thefluorescence emitter. Expanding the capability of a combinedSFM/FM set up to quantitative analysis of fluorescent signals islikely achievable based on characterizing calibration objects, suchas defined identifiable fluorophores, to be included within thesample.

5. Conclusions

The combined SFM/fluorescent microscope will allow identi-fication of specific proteins within complex assemblies on DNAand other biologically relevant molecular complexes. Appropriatemarker objects are essential to achieve nanometer accuracy inalignment of optical and topographic images when the moleculesof interest are smaller than optical resolution. We describe herethe convenient use of fluorospheres as robust fiduciary markers.In addition, using an optically suitable mica glued glass samplesubstrate, we take advantage of this atomically flat surfacecharacterized in greater than 15 years of SFM publications andwell established protocols for deposition and characterization ofbiological molecules like DNA–protein complexes. Our testsample, the RAD51 filament protein–DNA complex, reveals theadditional structural information that can be obtained from nmresolution topography images of objects identified by thepresence of fluorescent proteins.

Acknowledgments

This work was supported by a Marie Curie grant from theEuropean Commission (to H.S.), a VICI (to C.W.) and a TOP (to R.K.)grant from the Netherlands Organization for Scientific Research(NWO),Chemical Sciences division the Netherlands GenomicsInitiative/NWO and NCI (USA) program project (CA92584).

Appendix A. Supplementary material

Supplementary data associated with this article can be foundin the online version at doi:10.1016/j.ultramic.2010.03.002.

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