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* Department of Biophysics, University of Pécs, Faculty of Medicine, Pécs H-7624, Hungary; Institute of Physical Chemistry, University of Heidelberg, 69120 Heidelberg, Germany; Supertech Ltd., Pécs H-7624, Hungary; and Asylum Research, Santa Barbara, California 93117
Correspondence: Address reprint requests to Miklós S. Z. Kellermayer, Tel.: 36-72-536-271; Fax: 36-72-536-261; E-mail: miklos.kellermayer.jr{at}aok.pte.hu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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). In the past decade AFM has been increasingly used in biological applications owing to improvements in sample preparation, development of different data acquisition modes (noncontact mode) and experimental geometries (e.g., force spectroscopy (2)), and the ability of functionalizing the AFM tip that interacts with the sample (3).
There have been numerous attempts to combine AFM with fluorescence imaging (4–6). The motivation for combining AFM with fluorescence arises from the desire to exploit the complementary advantages of both techniques. Whereas AFM provides high spatial resolution and the possibility of mechanical manipulation, fluorescence offers high temporal resolution, sensitivity to local physical chemistry, and the possibility of functional imaging via specific labeling procedures. Imaging with combinations of AFM and various fluorescence microscopies (epifluorescence, confocal, total internal reflection fluorescence (TIRF)) have most often been sequential rather than simultaneous, owing to the difficulties in geometric arrangement and grossly different acquisition timescales. TIRF microscopy (TIRFM) (7), due to the narrow excitation field and its ability to detect and follow individual fluorophores, may offer special advantages in AFM-fluorescence combinations. Recently, individual green fluorescent protein (GFP) molecules picked up by an AFM cantilever tip were imaged with TIRFM (8), and simultaneous force and fluorescence measurements were achieved by following the emission of a fluorescent microsphere glued to a cantilever tip and excited with a calibrated evanescent field (9).
In this work, we developed a spatially and temporally synchronized TIRF (STIRF) and AFM system. The instrument was validated and tested on various cellular and biomolecular systems. With the technique, it is possible to correlate topography and fluorescence features of the examined specimen to mechanically manipulate soft biomolecular samples in a targeted fashion and to follow changes in the fluorescence properties of mechanically stretched biomolecules with high time resolution.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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), myosin (11), heavy meromyosin (HMM) (12), and titin (13) were purified according to established methods. F-actin was fluorescently labeled with molar excess of tetramethylrhodamine-isothiocyanate-phalloidin (TRITC-phalloidin, Sigma, St. Louis, MO). Titin was labeled with 
100x molar excess of tetramethylrhodamine iodoacetamide (TMRIA) or Oregon Green 488 maleimide (Molecular Probes, Eugene, OR) (14). Unbound fluorophores were removed by gel filtration (CL-2B Sepharose).
Actin-filament imaging
To image actin, fluorescently labeled filaments were immobilized on a surface coated with HMM (12,13,15). Briefly, a 24 x 60-mm glass coverslip (No. 1) was coated with 1% nitrocellulose (in isoamylacetate, Fullam, Latham, NY) in a home-built spin processor. Two parafilm stripes, forming spacers, were positioned in parallel on the coverslip, and an 18 x 18-mm glass coverslip was added on top so that a flow-through microchamber suitable for TIRFM imaging was constructed. A 10-µl aliquot of HMM (100 µg/ml) diluted in assay buffer (AB, 25 mM imidazole-HCl, pH 7.4, 4 mM MgCl2, 2 mM EGTA, 25 mM KCl, 1 mM dithiothreitol (DTT)) was first added to the chamber. Subsequently, the surface was blocked with bovine serum albumin (BSA, 0.5 mg/ml in AB). TRITC-phalloidin-labeled actin filaments (7-nM final concentration) were then added. To reduce photobleaching, the final buffer contained 100 mM ß-mercaptoethanol and an oxygen-scavenger enzyme system (16). To access actin filaments with the AFM cantilever tip, the top coverslip was gently removed from the flow-through microchamber.
Cell culture
HeLa cells were a kind gift from the Immunology and Biotechnology Department, University of Pécs, Faculty of Medicine. Human pancreatic cancer cells (Panc-1) expressing K8 and K18 keratins were a kind gift from the Medical University of Ulm, Germany. Cells were maintained in Dulbecco's modified Eagle medium (DMEM) containing 10% fetal bovine serum, 2% L-glutamine, 1% penicillin-streptomycin at 37°C in a humidified 5% CO2 atmosphere. For imaging, the cells were passed onto round coverslips (25-mm diameter, No. 1) and grown to subconfluence. Before experiments, Panc-1 cells were washed twice in serum-free DMEM and subsequently incubated for 18 h in serum-free DMEM.
Fluorescent labeling of cytoskeletal filaments
Washing and staining of samples were carried out in six-well plates. Cells were washed with phosphate-buffered saline solution (PBS, 10 mM K-phosphate, 140 mM NaCl), then permeabilized with Triton X-100 (0.1% in PBS) for 30 min at room temperature. Cells were fixed with paraformaldehyde (2% in PBS) for 15 min, washed with ethanolamine (10 mM in PBS) for 5 min, then rinsed with PBS. The actin-filament cytoskeleton was labeled with TRITC-phalloidin (200 nM in PBS complemented with 1% BSA) for 20 min at room temperature. Unbound TRITC-phalloidin was removed by washing with PBS. In the case of Panc-1 cells, keratin intermediate filaments were labeled with anti-pan-cytokeratin antibodies (KL1, Beckman-Coulter, Palo Alto, CA) for 30 min at room temperature, then the samples were labeled with fluorescein-isothiocyanate (FITC)-conjugated anti-mouse IgG antibodies. The round coverslips carrying the cells were mounted on a thin (2-mm) large-bore (23-mm) glass ring plate with silicon grease, so that the sample could be positioned on the microscope stage and accessed with AFM cantilever tip and objective lens at the same time. The cells were left covered with PBS complemented with 100 mM ß-mercaptoethanol to reduce photobleaching.
Synchronized TIRF/AFM Imaging
Fluorescently labeled cells and molecules were visualized with a partly custom-built STIRF and AFM combination (Fig. 1). The objective-based TIRF microscope is built on an Olympus IX81 motorized epifluorescence microscope with a TIRF module (Olympus, Tokyo, Japan) and a 60x, 1.45 numerical aperture oil-immersion objective lens. Light from a green laser (YAG 532 nm, 50 mW, JDS Uniphase, Milpitas, CA) and a blue laser (488 nm, 15 mW, Novalux, Sunnyvale, CA) were joined with a long-pass dichroic mirror and coupled to the TIRF module via an optical fiber (Point Source, Hamble, UK). The angle of incidence was adjusted by positioning the free fiber end in the plane conjugate to the back focal plane of the objective lens.
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300 kHz) or in liquid (BioLever, lever B, Au-coated on both sides, resonance frequency 35 kHz). Images (512 x 512 pixels or 1024 x 1024 pixels) were collected at a typical scanning frequency of 1 Hz. To obtain temporal synchrony, the fluorescence and AFM signals were simultaneously recorded and stored in parallel data sets ("waves", IgorPro, Wavemetrics, Lake Oswego, OR).
Focused TIRF microscopy
To reduce the size of the illuminated area to the diffraction limit and to increase excitation intensity, we used a focused TIRF arrangement (Fig. 1, lower inset). This arrangement is analogous to a confocal microscope in which a stationary, but tilted (and focused) laser beam is used for illumination. The instrumental arrangement was identical to that of wide-field TIRFM with one exception: we used a fiber optic cable with a collimator to couple the laser beam into the TIRF unit of the microscope. Thus, a collimated beam entered the back aperture of the objective, in contrast to the focused beam used in wide-field TIRFM. Tilting of the illuminating beam was accomplished by positioning the fiber-coupling assembly in the conjugate plane of the objective back aperture. The relatively large diameter (2 mm) of the beam entering the objective back aperture resulted in a wide range of incident angles at the sample plane (Fig. 1, lower inset). Only in the case of the largest incident angles did we obtain total internal reflection. Rays arriving at the sample at angles smaller than the critical angle became refracted. Thus, the excitation beam was partitioned into a reflected and a refracted portion. In our experiments we positioned the excitation laser beam in such a way that the rays passing through the sample were tilted away from the cantilever, but the beam was entirely within the back aperture.
Mechanical manipulation of fluorescently labeled, surface-attached soft samples
Mechanical manipulation of TRITC-labeled, cultured HeLa cells or individual, HMM-attached actin filaments was carried out by first coaligning, to within 1 µm, the AFM cantilever (Olympus AC160TS) with the optical axis and scanning an initial TIRF microscopic image. Subsequently a user-defined nanolithographic pattern was drawn into the surface with a typical pressing force of 1 nN, using the MicroAngelo program routines of the AFM driving software (Asylum Research). Finally, a second scanning TIRFM image was obtained.
Single-molecule force spectroscopy of fluorescently labeled titin
TMRIA-labeled titin molecules were first attached to the AFM cantilever (Olympus BioLever, lever B) by pipetting an aliquot (5 µl) of sample (0.05 mg/ml) onto the cantilever tip and incubating at room temperature for 10 min. Unbound molecules were removed by successive washes with buffer (AB with 200 mM KCl). The titin-coated cantilever was pressed in the evanescent field of wide-field TIRFM and brought near a cleaned microscope coverslip. The cantilever tip was coaligned with the optical axis by moving the tip in the XY direction and seeking the maximum in the APD signal. The cantilever tip was than pressed onto the coverslip surface with constant rate (typical rates 500 nm/s). Cantilever displacement and deflection signals were collected. Cantilever stiffness was calibrated with the thermal method (17–19). Typical cantilever stiffness was 30 pN/nm.
Data analysis and processing
Image processing was carried out using the built-in routines of the MFP3D software. Three-dimensional (3D) surface rendering was done with the Argyle toolbox of the MFP3D software. For calculations and statistical analyses we used IgorPro (v. 5.0) and KaleidaGraph (v. 4.0) software packages. Whenever necessary, nonlinear force curves were fitted with the worm-like chain model of entropic elasticity (20).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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Stage-scanning, evanescent field excitation confocal microscopy
For fluorescence imaging in synchrony with AFM data acquisition, we first developed a stage-scanning, evanescent field excitation confocal microscope (Fig. 1). In this arrangement the fluorescence image was obtained by scanning the sample in the evanescent field and collecting photon count or intensity data at each point of the scan. The size of the illuminated area was limited by using a field diaphragm positioned in the excitation beam path in a focal plane conjugate with the sample. To maintain high spatial resolution, a pinhole (40 µm) was positioned in front of the photon counting module. The signal from the photon counting module was either used directly as number of photons per image pixel or converted to intensity by using a rate meter. The pixel size, set by the ratio of the scan size (5–70 µm for the images herein) and image resolution (typically 512 x 512), varied between 10–140 nm. In terms of image quality we found no significant difference between the two acquisition methods.
We tested the stage-scanning, evanescent field excitation confocal microscope on samples of surface-attached actin filaments and HeLa cells in which the actin cytoskeleton was labeled with TRITC-phalloidin (Fig. 2). The fluorescence image of TRITC-labeled actin filaments attached in rigor state to a HMM-coated surface is shown in Fig. 2 a. The image is essentially indistinguishable from that recorded with a microchannel-plate-intensified charge-coupled device (CCD) camera (data not shown). The half-maximum width of the Gaussian intensity profile across the actin-filament image (Fig. 2 b) was 394 nm, which is comparable with the optical resolution for this emission wavelength (580 nm). We found that the root mean-square (RMS) variation of the fluorescence intensity along an actin filament in the evanescent field was 60 in contrast to that of the background, which was 17 (Fig. 2 c). The large spatial variation of emission intensity along the actin filament might be due to a variation of the filament position in the evanescent field. The scanning TIRF image of TRITC-phalloidin-labeled HeLa cells is shown in Fig. 2 d. Bundles of actin filaments running parallel to the microscope coverslip surface can be clearly visualized.
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300 nm apparent between the AFM (Fig. 4 d) and the TIRFM (Fig. 4 f) images also indicates that a residual misalignment is present between the cantilever tip and the optical axis.
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) are observed.
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Mechanical manipulation with STIRF-AFM
Observations on the mechanical manipulation of cells and molecules with STIRF-AFM are shown in Fig. 8. In the first set of experiments we used STIRF-AFM to ablate parts of surface-attached cells and actin filaments (Fig. 8, a and b). Because of spatial synchrony, areas to be mechanically manipulated can be defined and targeted with high (submicron) precision. As evidenced by the images, targeted parts of the surface-attached HeLa cells and actin filaments were successfully removed with the technique.
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,22), were observed. In the fluorescence traces sometimes we observed abrupt increases in fluorescence intensity, indicating that a structural change probably occurred in the part of the titin molecule that remained in the evanescent field. In the case of tip-adsorbed titin, molecules were stretched by lowering the tip onto the coverslip surface, then pulling the tip away with a given stretch velocity (Fig. 8 d, inset).
If titin molecules became attached to the coverslip surface, then these molecules became stretched by the procedure. In these experiments the fluorescence traces were dominated by a single-exponential decay that most likely corresponds to the movement of the fluorescent tip in the exponentially decaying evanescent field (Fig. 8 d). In support of this notion, the movement of uncoated Si3N4 tip, which we found to be brightly fluorescent at the 532-nm excitation wavelength, also resulted in exponentially decaying fluorescence intensity (Fig. 8 e). By decreasing the angle of incidence of the excitation laser beam, the depth of the evanescent field increased as previously observed (9). In the case of tip-adsorbed titin, we also observed sudden changes in fluorescence intensity (Fig. 8 d, asterisk).
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DISCUSSION |
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There are numerous motivations for combining AFM with fluorescence. AFM provides high-resolution topography and mechanics (viscoelasticity) information of the examined specimen (1). Furthermore, it is possible to directly manipulate the sample mechanically (22,23), a method that has evolved into single-molecule force spectroscopy (2). Various modalities of fluorescence (intensity, polarization, anisotropy, and lifetime) provide dynamic and structural information related to the local physical-chemistry of the specimen (3,24,25). In certain fluorescence modes (fluorescence resonance energy transfer, FRET) structural information of very high resolution may be obtained (26,27).
There have been many attempts to combine AFM and fluorescence imaging (4–6,8,9,28,29). Due to the vast differences in acquisition rates and modes of imaging, the acquisition of AFM and fluorescence has usually been sequential. A direct correlation of the fluorescence and AFM data in these methods is difficult, and typically they are segregated in separate data sets. To establish fully synchronized AFM and fluorescence imaging, we carried out sample scanning and coaligned the optical and mechanical axes. In this imaging mode, at each point of the sample scan the mechanical (cantilever deflection) and fluorescence (emitted photons) information were recorded simultaneously.
As a first step in establishing synchronized fluorescence and AFM imaging, we developed a sample-scanning fluorescence imaging method. Ideally, fluorescence excitation should be spatially limited to the part of the sample being examined with the AFM cantilever tip, but the excitation beam should not interfere with the cantilever itself. We experimented with two excitation geometries, each of which partly fulfills the ideal requirements. We used TIRF, in which the depth of the evanescent field that excites the fluorophores is on the order of a couple hundred nanometers (7), and therefore interference with AFM imaging is unlikely since the cantilever tip is typically >10x longer than this. First, we used wide-field, objective-based TIRF microscopy (Figs. 1, upper inset, 2–6
, and 8, c–e), in which an illuminated area of the sample was limited with a field diaphragm to 5 µm. Second, we used a focused TIRF arrangement, in which the sample was illuminated with a focused and tilted laser beam (Figs. 7 and 8, a and b).
Focused TIRFM in an arrangement resembling dark-field illumination was recently used to excite attoliter volumes of fluorescently labeled sample (30). In our arrangement of focused TIRFM, only part of the participating rays is totally reflected, because they arrive at the sample at a wide range of incident angles. Nevertheless, the refracted rays do not interfere with the cantilever either. We found that very high signal/noise ratio and low photobleaching are obtained with scanning, focused TIRFM. In the case of stationary samples used in single-molecule force spectroscopy (Fig. 8, c–e), however, wide-field TIRFM may be better suited. The advantages of focused TIRFM over the wide-field arrangement are 1), high excitation intensity, 2), low photobleaching, and 3), improved spatial resolution (alleviation of the need for a confocal pinhole). The main disadvantage of focused TIRFM is its cumbersome alignment: the excitation focal point must be coaligned with the detection volume for an in-focus specimen by tilting the laser beam in the conjugate focal plane of the objective back aperture.
The spatial resolution reached with scanning TIRFM is on the order of the diffraction limit (Fig. 2). In fact, images obtained on single actin filaments with scanning TIRFM are indistinguishable from those obtained with microchannel-plate-intensified CCD cameras. Furthermore, because of the narrow excitation field, local variations in fluorescence intensity, possibly related to filament position relative to the surface, may be detected (Fig. 2 c). The time resolution of scanning TIRFM imaging matches that of AFM scanning. Therefore, obtaining a TIRFM image takes several minutes. Because of the low image acquisition rate and the narrow excitation field, focusing is often difficult. Refocusing during scanning requires precautions because movement of the oil-immersion objective, which is in viscous coupling with the sample, causes noise in simultaneous AFM scanning. We are currently developing means to establish automatic focusing that addresses the problem of the additional mechanical noise. In contrast to the low image acquisition time resolution, when fluorescence is measured at a single, user-defined point, acquisition rates up to 100 kHz (limited by analog-to-digital conversion rate) may be reached.
We successfully combined scanning TIRFM imaging with AFM (Figs. 3–7). Complete temporal synchrony was achieved because of simultaneous acquisition of the photon count and AFM data. Spatial synchrony was obtained by positioning the cantilever tip onto the optical axis. That is, the cantilever tip was positioned on the location sensed by the photon-counting module. In this work, we established spatial synchronization by positioning, under video control and using stage micrometers, the cantilever tip to an empirically determined location before the scanning experiment. Most often we were able to achieve spatial synchrony within the diffraction limit (see Fig. 5). At higher magnifications, however, residual discrepancies between spatial correspondence became evident (Fig. 4). Furthermore, spatial synchronization is sensitive to drift, and constant readjustments, using closed-loop piezo translators and fiducial image coordinates, may be necessary. Using STIRF-AFM we obtained spatially and temporally synchronized images of monolayer cells in culture.
To validate the method, in this work we used fixed and dried cells, which are easier to scan with AFM. Microfilament (Figs. 3, 4, and 6) and intermediate filament (Fig. 6) components of the cytoskeleton could be visualized and localized with the method and matched with surface topography elements. Importantly, because of synchronized AFM and optical imaging, problems associated with independent nonlinearities of the two imaging methods are alleviated. We successfully employed STIRF-AFM to image surface-adsorbed, fluorescently labeled titin molecules (Fig. 5). Optical resolution issues related to the diffraction limit become particularly visible with our imaging method, because single fluorescent spots often contained two or more titin molecules as revealed with the AFM scanning (Fig. 5 a). STIRF-AFM may aid in identifying the exact number of molecules behind a single spot in the fluorescence image.
AFM tip fluorescence may interfere with obtaining high-quality fluorescence images by reducing signal/noise ratio. Tip fluorescence emission depends on the cantilever type, excitation wavelength, and excitation intensity. We found that uncoated silicon nitride tips fluoresce when excited with 532 nm, but emission intensity is significantly reduced at 488-nm excitation. Gold coating on the tip surface significantly reduced tip fluorescence, probably because much of the excitation intensity was reflected by the tip.
A special application of STIRF-AFM is the mechanical manipulation of fluorescently labeled cellular and biomolecular samples (Fig. 8). Because of spatial synchrony, it is possible to first image the specimen with scanning TIRFM, then manipulate the sample at user-defined locations with AFM using nanolithography protocols, and finally reimage the sample with TIRFM to detect the evoked changes. We have successfully applied this method to ablate parts of cells and individual actin filaments (Fig. 8, a and b). A particularly interesting application of STIRF-AFM is in the exploration of mechanically driven structural changes in biomolecules. Currently very little is known about the details of structural changes evoked in single molecules by mechanical manipulation. Structural knowledge is inferred from particular aspects of the mechanical data (e.g., contour length calculated from fitting with polymer models) compared with independent structural data (e.g., contour length calculated from protein sequence) (22,31,32). A simultaneous but independent method that provides structural information may reveal the details of intramolecular changes evoked by mechanical perturbation. In the TMRIA-labeled titin-molecule strand stretched with AFM we sometimes observed sudden increases of fluorescence intensity (Fig. 8, c and d).
These bursts of fluorescence intensity were seen both during mechanical stretch and relaxation. We have previously shown that the fluorescence of TMRIA-labeled titin is quenched in the native state of the protein due to the formation of rhodamine dimers (14). Indeed, the 518-nm peak in the absorption spectrum of TMRIA-labeled titin used here (Fig. 8, f) revealed the presence of rhodamine dimers. Fluorescence intensity increases upon the addition of chemical denaturants to TMRIA-labeled titin (14). Possibly, the mechanical extension of titin and the concomitant unfolding of the molecule resulted in the dissociation of rhodamine dimers and hence in the increase of fluorescence intensity observed here. So far we have not been able to correlate the observed fluorescence changes with particular mechanical events such as the sawtooth-shaped force transitions. Furthermore, the detected fluorescence changes originate from parts of the titin molecule that remain in the evanescent field during the mechanical manipulation. It is also possible that sudden changes in the concentration of fluorophores within the excitation volume, caused by relaxation and contraction of the titin molecule, may also contribute to the observed fluorescence changes. Further work will help to sort out the exact mechanisms of our findings.
Further applications, widespread use, and future development of the STIRF-AFM instrument will probably depend on the ease of assembly and use, the costs involved, and the possibility of combination with other novel microscopic techniques (e.g., single-molecule polarization methods (33), FIONA (34)). The costs involved in constructing the instrument herein, in addition to the AFM and the light microscope, are related to lasers, coupling optical fibers, APD detector, and an intensified CCD camera for alignment purposes (altogether $30,000). Although setting the instrument up is relatively easy, frequent realignments are necessary to achieve spatial synchrony and high-quality images. Coalignment and autofocus algorithms, currently under development, may aid in improving the ease of use. Finally, the use of high-speed, high-resolution cameras instead of APD may expand the applicability of STIRF-AFM to novel fluorescence imaging methodologies.
In sum, a synchronized TIRF and AFM was developed and tested. The instrument, which we named STIRF-AFM, allows the visualization of surface topography and fluorescence of suitable cellular and biomolecular samples simultaneously. Furthermore, STIRF-AFM makes it possible to follow the fluorescence of mechanically manipulated molecules with high time resolution. Conceivably, STIRF-AFM may contribute to resolve the details of structural changes that occur in biological molecules upon mechanical manipulation.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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Submitted on March 20, 2006; accepted for publication July 6, 2006.
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REFERENCES |
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