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Abstract
Real-time optical imaging combined with single-molecule manipulation broadens the horizons for acquiring information about the spatiotemporal localization and the mechanical details of target molecules. To obtain an optical signal outside the focal plane without unintended interruption of the force signal in single-molecule optical imaging-force spectroscopy, we developed an optical method to extend the depth of field in a high numerical aperture objective (≥ 1.2), required to visualize a single fluorophore. By axial scanning, using an electrically tunable lens with a fixed sample, we were successfully able to visualize the epidermal growth factor receptor (EGFR) moving along the three-dimensionally elongated filamentous actin bundles connecting cells (intercellular nanotube), while another EGFR on the intercellular nanotube was trapped by optical tweezers in living cells. Our approach is simple, fast and inexpensive, but it is powerful for imaging target molecules axially in single-molecule optical imaging-force spectroscopy.
© 2017 Optical Society of America under the terms of the OSA Open Access Publishing Agreement
1. Introduction
Single-molecule force spectroscopy, including optical tweezers, magnetic tweezers, and atomic force microscopy (AFM), is highly suitable for the investigation of molecular-level forces and mechanical properties of biomolecules or cells under applied forces [1]. Force spectroscopy combined with real-time optical imaging provides multi-dimensional information on biological structures and functions of biomolecules in vitro or in cells [2–5]. The most common optical imaging of a probe associated with biological molecules is fluorescence microscopy. Both total internal reflection fluorescence (TIRF) microscopy and confocal microscopy have been adopted to achieve a particularly high signal-to-noise ratio (SNR), which is required for single-molecule fluorescence imaging in single-molecule force spectroscopy-fluorescence microscopy (smFS-FM) [4–6]. However, TIRF microscopy is not suitable for the visualization of probes that span several micrometers along the axial direction because the evanescent wave generated in the TIRF microscopy only penetrates the sample by ~200 nm. A conventional method to visualize a single probe moving along the axial direction is to use a confocal microscope with a piezoelectric sample stage that can axially scan to detect the probe in the axial direction, with the focal plane fixed. However, the vertical movement of the sample stage interrupts the force sensor linked to a target biopolymer, which is fixed to the surface, resulting in an unintended interference with the force measurement. Another method is to control the position of the objective instead of the sample stage piezo-electrically [7], however it is not applicable to single-molecule fluorescence imaging using an objective with high magnification and high numerical aperture (NA). It is also not suitable to smFS–FM in an optical trap through an objective, because the focal plane of the trapping laser is varying with the objective moves. Thus, this technical problem limits the combined applications of force spectroscopy methods, especially magnetic tweezers and AFM, with fluorescence imaging.
In this study, we introduce an application of an electrically tunable lens (ETL) to visualize the axial motion of a fluorescent probe in optical tweezers, combined with single-molecule fluorescence microscopy. The ETL tunes its focal length with a quick response to the input voltage or current, which facilitates the fast switching of the focal plane at tens of μm-spatial and ms-temporal resolution, without the mechanical movement of the imaging system. However, optical properties, such as magnification and focal shift, can change at different ETL focal lengths over a wide axial range, which demands complex optical modifications to overcome these problems [8, 9]. The ETL was used in cells of deep tissue imaging by scanning frame-by-frame along the axial (z-axis) direction, however it has not yet been used for single-molecule imaging [10, 11]. In single-molecule confocal microscopy incorporated with optical tweezers, we effectively extended the depth of field of the confocal microscope by fast axial scanning of the ETL in a frame in a high NA objective lens (NA = 1.2 and 1.4) without moving the sample stage or the objective in the axial direction. We demonstrated the performance of the ETL system in the single particle tracking analysis of epidermal growth factor receptor (EGFR) on filamentous actin bundles connecting cells (intercellular nanotube), in the presence of the mechanical force applied to the intercellular nanotube, using an optical trap combined with a confocal microscope.
2. Materials and methods
2. 1 Optical tweezers combined with a line-scan confocal microscope
We constructed a video-rate line-scan confocal microscopy (LSCM) device [12] based on commercial optical tweezers (NanoTracker 2, JPK Instruments AG) (LSCM–OT) in an inverted microscope, as shown in Fig. 1. Details of the optical tweezers system are described on the company homepage (www.jpk.com/products/force-sensing-optical-tweezers-and-optical-trapping/nanotracker-2). A detector objective lens (LUMPlanFLN 60 × , NA = 1.0, water dipping, Olympus) was used for collecting light scattered from a force sensor (a polystyrene bead of 2 μm or a fluorescent polystyrene bead of 1 μm in diameter). A high NA objective lens (UPlanSApo 100 × , NA = 1.4, oil immersion or 60 × , NA = 1.2, water immersion, Olympus) focused the fluorophore-excitation (532 nm) and bead-trapping (1064 nm) laser beams. Fluorescent emission signals were acquired through the trapping objective lens (Fig. 1). To avoid a significant change in magnification while focusing by an additional ETL in the confocal microscopy pathway, an ETL (EL-10-30-C, Optotune) was inserted between two telescopes (T1 with f = 400 mm pair, T2 with f = 200 mm pair), where the conjugated plane of the back focal plane of the trapping objective lens in the excitation-emission path of the LSCM was located. The ETL was placed on a plane parallel to the sample stage to prevent asymmetrical deformation of the ETL polymer membrane due to gravity. Two dichroic beam splitters (DM1: 776 long-pass / DM2: Di01-R405/488/532/635, Semrock), another telescope (T3 with f = 100 mm pair), two Galvano scanning mirrors for line confocal scanning (6231H, 15 mm, Cambridge Technology), a 100 μm slit (S100R, Thorlab), and a band pass filter (FF01-515/588/700-25, Semrock) were aligned in the line-scan confocal microscopy (Fig. 1). The fluorescence signal was imaged using an electron multiplying CCD (EMCCD) camera (iXon3, Andor).
Fig. 1 Schematic of the line-scan confocal microscopy (LSCM) combined with optical tweezers (OT). After passing through the cylindrical lens (CL), the shape of the excitation beam changes from an enlarged circle to a thin oval. The box at the bottom displays cross sections perpendicular to the propagation of the excitation beam at each position of a, b, c, and d in the path of the solid (x-axis view) and dashed (y-axis view) lines. The red solid line is the path of the emission signal. The grey solid line is the path of the trapping laser and the optical signal from the trapped bead. GM: Galvano scanning mirror, DM: Dichroic mirror, BX: Beam expander, T1/T2/T3: Telescope. The 3D detection and Trapping system is the part of the NanoTracker2 from JPK Instruments AG.
2.2 Living cell experiment
A polydimethylsiloxane (PDMS) flow chamber (25.0 mm × 3.0 mm × 0.7 mm) was constructed for the efficient addition and washing of ligands and fluorophores in live cell experiments. HeLa cells (passage number > 3) grown in 10% fetal bovine serum (FBS) and phenol-red free Dulbecco’s modified eagle medium (DMEM) (Capricorn) were flowed in the PDMS chamber by using a syringe pump to achieve 70-80% confluence, then incubated overnight at 37°C. In addition, the heating sample stage of the microscope was applied to maintain a sample temperature of 37°C (Live Cell Instrument). Fluorescent quantum dot-coated epidermal growth factor (EGF–QD) was prepared at a ratio of 5:1 with biotin-conjugated EGF (Molecular Probes) and streptavidin-conjugated quantum dot 605 (Invitrogen). We also prepared EGF–coated beads (EGF-bead) with streptavidin-coated, fluorescent polystyrene microsphere (diameter of 1 μm, Bangs Laboratory) and biotin-conjugated EGF at a ratio of 1:100,000. HeLa cells were starved for more than 1 hr in FBS free media and incubated in DMEM with 1 nM EGF–QD for 15 minutes. After free EGF–QDs were washed, 5 fM EGF-beads in serum-free DMEM were added. The optical tweezers trapped an EGF-bead, bound to the intercellular nanotube, and the LSCM visualized fluorescent signals from EGF–QDs on the intercellular nanotube. All optical trap controls, including laser power adjustment, force calibration by power spectrum measurement, real-time force signal measurement, and force clamping, were managed by the JPK NanoTracker Desktop Software (JPK Instrument AG). The trap stiffness of the EGF-bead was calibrated as kx = 0.253 ± 0.014 pN/nm in the x-direction and ky = 0.258 ± 0.058 pN/nm in the y-direction. The force and fluorescence signals in the trapping experiments were obtained for every 100 Hz and 10 Hz, respectively. A force clamp was employed using a feedback system that measures the instantaneous position of the trapped bead, then moves the trap along the longitudinal axis of the intercellular nanotube at a rate of 2 μm/s, to maintain the displacement set between the bead and the trap center at 10 Hz [13]. We analyzed the data using the JPK Data Processing software (JPK Instrument AG) and a custom-built Matlab program.
3. Results and discussion
3.1 ETL setup with no significant signal loss and with a fixed magnification
The ETL and Galvano scanning mirrors (GM1 and GM2) in the confocal path of the LSCM–OT were synchronized [Figs. 1 and 2(a)]. The input voltage was set as a triangular waveform with an amplitude of Vscan [Fig. 2(a)]. The ETL scanning frequency was identical to the sampling rate of the EMCCD (10 Hz), while the Galvano scanning mirrors scanned the imaging plane at the frequency of 50 Hz. The ETL system consists of an ETL and an offset lens with a negative focal length (f = −100 mm). As the offset voltage (V0) was set to 3 V, the ETL system had a negative focal length for input voltages less than 3 V (VETL < 3 V) and a positive focal length for input voltages greater than 3 V (VETL > 3 V) [Fig. 2(b)]. To find the optimal single-molecule imaging conditions, we quantified the optical properties of the LSCM–OT over the focal length of the ETL using a fluorescent bead of 60 nm in diameter (Bangs Laboratory) coupled on a piezo-electrically controlled surface. The object was viewed about 1.1 times the magnification of the trapping objective lens, but it was independent of the focal length of ETL (VETL) [Fig. 2(c)]. Nevertheless, the SNR of the object varies with VETL: it apparently decreases as VETL increases or decreases from the offset voltage and it is reduced by 62% at VETL = 4 V in a 100x objective lens [Fig. 2(d)]. The SNR was determined by , where is the intensity of the object, is the intensity of the background, and is the standard deviation of the background intensity. Nonetheless, the SNR was maintained with no significant loss in the range of ± 0.5 V from the offset. The focus shift appears to be in a linear region of a slope of −10.2 ± 0.1 μm/V (100 × objective lens) or −20.7 ± 0.3 μm/V (60 × objective lens) at the input voltage range from 2 V to 4 V [Fig. 2(e)]. Considering these, by tuning the ETL between an input voltage range from 2.5 V to 3.5 V we can focus on probes in a span of up to 10 μm along the axial direction in a 100 × objective lens (or 20 μm in a 60 × objective lens) with a fixed magnification and without any significant signal loss.
Fig. 2 (a) Input voltage control of Scan OFF and ON with a 10 Hz ETL using a triangle wave. The Galvano scanning mirror scans at a frequency of 50 Hz and the sampling rate of the EMCCD is 10 Hz. (b) The offset lens in front of the ETL adjusts the focal length of the ETL to infinity for VETL (=V0) = 3 V, negative for VETL < 3 V, and positive for VETL > 3 V. (c) Magnification of the LSCM in a 100× or 60× objective lens as a function of the focal length of the ETL (VETL). (d) Relative SNR with respect to SNR at VETL = 3.0 V at various VETL. (e) Focal shift in the microscope with a 100× or 60× objective lens at various VETL
3.2 Fluorescence imaging along the axial direction
Figure 3 shows the depth of field measurement using a long double stranded DNA (dsDNA) molecule, which was stained by the SYTOX Orange intercalator (ThermoFisher Scientific) in a 100 × oil immersion trapping objective lens. A 21.5-kb DNA molecule was constructed as previously described [14]. The 5′-biotinylated end of the 21.5-kb DNA was attached to a biotin-polyethylene glycol-coated coverslip using a biotin-streptavidin interaction, and the opposite 3′-digoxigenin (dig) end was linked to a polystyrene bead (diameter of 2.0 μm) functionalized with an anti-dig antibody (Spherotech) [Fig. 3(a)]. A bead attached to a tethered DNA molecule was selectively trapped and then pulled by moving a piezo stage along the x-direction with a trap stiffness of kx = 0.141 ± 0.012 and kz = 0.038 ± 0.009 pN/nm, causing the center of the bead to be 6 μm above the surface and the DNA molecule to stretch by ~7.0 μm [the crystallographic length = 21.5 × 103 bp × 0.34 nm/bp = 7.3 μm; Fig. 3(a)]. Only a limited portion of the fluorescently stained DNA appeared to be visible and the focal plane was moving down from the bead when VETL increases from VETL = 3.0 V to 3.6 V with no scanning (Scan OFF) [Figs. 2(a) and 3(a)]. However, fast scans of VETL = 3.0 V to 3.6 V (or 3.1 V to 3.5 V) within a 100 ms time resolution (Scan ON) clearly extend the visible range to nearly all DNA length [Fig. 3(b)]. The emission intensity profile of dsDNA stained with Sytox Orange in Scan OFF obtained from the kymograph (VETL = 3.3 V, gray solid) and in Scan ON (VETL = 3.3 ± 0.3 V, magenta solid) can be seen in Figs. 3(a) and 3(b), respectively [Fig. 3(c)]. We determined the depth of field by measuring the full width at half maximum (FWHM) of the intensity profile [Fig. 3(c)] using the relation of Depth of field = FWHM*6.0/5.3 [see Fig. 3(a), drawing]. The intensity profile in Scan ON (VETL = 3.3 ± 0.3 V) is nearly identical to the average of the intensity at each VETL ( = 3.0 ~3.6 V) with a step of 0.1 V (magenta dot). The depth of field (1.6 ± 0.2 μm) at VETL = 3.0 V is significantly extended by 1.6 folds (2.6 ± 0.3 μm) and 2.2 folds (3.5 ± 0.2 μm) at 3.3 ± 0.2 V and 3.3 ± 0.3 V, respectively [Fig. 3(d)]. In this study, by using a 100 × objective lens, we demonstrated that the rapid change of the focal plane within the sampling rate (100 ms) was suitable to visualize probes that are within 3.5 μm in an axial direction in a 100 × objective with NA = 1.4 without signal loss and the interference of the trapped bead. The depth of field definitely increases in a trapping objective lens with lower magnification and NA (Fig. 2).
Fig. 3 (a) Sytox Orange-stained dsDNA immobilized on the surface pulled by optical tweezers (left). The stained DNA is imaged at a specific focal length of the ETL (right). (b) VETL varies from 3.0 V to 3.6 V or from 3.1 V to 3.5 V at a frequency of 10 Hz. The depth of field increases (right). (c) The intensity profiles of the stained DNA are presented in Scan OFF (gray), ON (magenta, solid), and the average of the intensities of the DNA at 3.0 V ~ 3.6 V in Scan OFF (magenta, dotted). (d) The depth of field is measured in Scanning OFF and Scan ON (3.3 ± 0.2 V and 3.3 ± 0.3 V).
3.3 EGFR mechanics on intercellular nanotube in live cells
This method was applied to extend the depth of field, to study the mechanics of the epidermal growth factor receptors (EGFRs) on intercellular nanotube connecting cells over a long distance (up to 100 μm). Extensive studies on the intercellular nanotube have revealed its biological significance in various transports of receptors [15–19], organelles [20], vesicles [21], virus [22, 23], morphogens [24–27], nucleic acids [28, 29] and mitochondria [30] in numerous cell types. EGFR is a key receptor of the signaling pathway for cell growth and proliferation, where activated-EGFR undergoes a systematic retrograde transport on filopodia consisting of actin filaments. The mechanics of actin filaments in a filopodium is linked to the transport of EGFR. Although it is critical to clarify the biological relevance of EGF signaling and the intercellular nanotube, the transport of activated-EGFR on an intercellular nanotube has still not been reported. The single-molecule tracking of EGFR might be the most direct and robust method to monitor the transport of EGFR on elongated intercellular nanotubes.
We visualized in real-time the individual EGF–QDs distributed on an intercellular nanotube between HeLa cells at a 100 ms time-resolution for 10 s [Fig. 4(a)]. Not all EGF–QDs on the intercellular nanotube were focused at a particular VETL. As VETL increased from 3.1 V to 3.5 V, the focused area apparently moved to the left, which indicates that the long intercellular nanotube is axially inclined between cells [see drawing in Fig. 4(b)]. In contrast, in Scan ON (VETL = 3.3 ± 0.3 V), the focused EGF–QDs were present in the lateral extent of the intercellular nanotube of 34 μm. The time-dependent position of the individual EGF–QDs shows that most EGF–QDs are not transported unidirectionally, but they move randomly along the intercellular nanotube with a diffusion coefficient of 1.2 ( ± 5.9) × 10−3 μm2/s [mean ± SD; Fig. 4(b)]. This result seems to be inconsistent with the dynamics of the activated-EGFR on filopodia: the EGF-bound EGFR requires a dimer formation for the activation of the downstream signaling [31], resulting in the unidirectional transport of the activated-EGFR on the filopodium, following the retrograde flow of actin filaments in the filopodium [32]. However, the addition of extra unlabeled-EGFs (3.4 nM) to the EGF–QD (1 nM) enables the observation of the unidirectional transport of EGFR on the intercellular nanotube, which is consistent with the previous observation on filopodia [Figs. 4(c) and 4(d)].
Fig. 4 (a) Schematic representation of EGFR imaging on the intercellular nanotube connecting HeLa cells. (b) Kymograph of the EGF–QDs on the intercellular nanotube with the focal shift of 4.1 μm (VETL = 3.1 ~3.5 V, step = 0.1 V). (c) Schematic of an EGFR transport using EGF coated bead as an additional force probe. (d) EGF–QD Kymographs, (e) trapping force, and (f) trap position for three different experiments.
Using the LSCM–OT, we monitored both the EGF–QD [red in color in Fig. 4(d)] and the EGF-bead [brighter and thicker lines in Fig. 4(d)] on the intercellular nanotubes to study the mechanism of the retrograde movement of the EGFR and the actin filaments in the intercellular nanotube. As expected, in the absence of the unlabeled EGF, the force acting on the trapped EGF-bead is apparently steadily zero in the x- and y-directions [Figs. 4(e) and 4(f), left] and most of the EGF–QDs diffuse along the intercellular nanotube [Fig. 4(d), left]. The addition of 3.4 nM EGF ligands in the solution causes the EGF–QDs move unidirectionally on the intercellular nanotube at a rate of 5.13 nm/s or 4.24 nm/s [Fig. 4(d), center], and the trapping force increases to 30 pN in the longitudinal direction of the intercellular nanotube [Fig. 4(d), center], due to the retrograde movement of the actin filaments. In force-clamping mode, the EGF–QD translocates unidirectionally at a rate of 14.5 nm/s [Fig. 4(d), right], while a constant force of 10 pN on the EGF-bead, moving at a rate of about 9 nm/s [Fig. 4(e), right]. This observation indicates that we successfully visualize the dynamics of a target molecule moving along the axial direction without interrupting the force acting on the system during the measurement.
4. Conclusions
We developed a fast axial scan system for single-molecule fluorescence imaging in a video-rate confocal microscope combined with optical tweezers, which extends the depth of field of a high numerical aperture objective lens (NA ≥ 1.2) without interruption of the trap. Using this system, we successfully visualized the transport of EGF receptors on the intercellular nanotube consisting of actin bundles between cells, while the EGF receptors are mechanically manipulated. The focal length of the ETL was adjusted with a scan voltage of 0.3 V, resulting in the doubling of the depth of field in the microscope. Although the depth of field can exhibit a length of 20 μm by adjusting VETL from 2 V to 4 V in a 100 × objective lens (NA = 1.4) [Fig. 2(d)], the scan voltage of the ETL has to be determined considering the magnification and SNR required for single-molecule imaging [Figs. 2(c), 2(d), and 2(e)]. In addition to cell applications using optical tweezers, the ETL-based axial scan system is well suited to smFS-FM experiments, such as magnetic tweezers and AFM, which manipulate biomolecules of DNA, RNA, or proteins in the vertical direction of the sample stage.
When we increased the amplitude of ETL scanning, the SNR was reduced due to the increase of the background signal by the extension of the scanning depth. However, there is a tradeoff between depth of field and SNR because the imaging depth expands at larger amplitudes. We also confirmed the ETL scanning performance by imaging a living cell labeled with EGF-QDs at different ETL scanning rates from the frame rate of the EMCCD (10 Hz). The same ETL scanning frequency as the video rate of EMCCD did not affect the depth of field. However, at an ETL scanning frequency higher than the video rate of EMCCD, we observed the degradation of images. Thus, the optimized amplitude of ETL scanning is the most critical variable in this system to obtain sufficient photon and SNR for single-molecule fluorescence imaging [see Fig. 2(d)]. The frequency of ETL scanning and the video rate of EMCCD should also be adjusted to collect enough photons from a single fluorophore and to obtain images without their deterioration.
In fact, ETL has been exploited in various microscopes, such as two-photon microscopy [9, 10], confocal microscopy [8, 11], phase microscopy [33], light-sheet microscopy [34], wide-field microscopy [35], photoacoustic microscopy [36], and HiLo microscopy [37]. These microscopes are intended to have a wide axial scan range of hundreds of micrometers in an objective lens with low NA (< 1.0) for deep-tissue imaging, except for one study [35], which used a high NA objective lens (NA = 1.4) in wide-field microscopy for the static image of protein-complexes in HeLa cells. Our study is a first demonstration of an ETL being used for a single-molecule force-optical imaging system in extended depth of field.
Funding
National Research Foundation (NRF) of Korea (2017K1A1A2013241, 2009-0081562).
Acknowledgments
We thank Yongmoon Jeon and Ryanggeun Lee for the help of the DNA construction. This research was supported by Global Research Lab Program through the National Research Foundation (NRF) of Korea funded by the Ministry of Science and ICT (NRF-2017K1A1A2013241 to J.-B.L.) and by Creative Research Initiatives (Physical Genetics Laboratory, 2009-0081562 to S.H.)
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