Using Combined Fluorescence Lifetime Imaging Microscopy and Atomic Force Microscopy to Perform Enhanced Multiparameter Cell Imaging

Table of Contents

Introduction
Benefits of Combined Techniques
     Live Cell Imaging
     High-Resolution Molecular-Scale Imaging
     Force and Mechanical Studies
Combined System Description
Synchronization of the Two Systems
Optical Alignment
Imaging of Fluorescent Beads
Fixed Cells
Conclusion

Introduction

Fluorescence lifetime imaging microscopy (FLIM) offers optically encoded data about live cells processes. In contrast, atomic force microscopy (AFM) provides mechanical and nanometer-resolved surface topography data, and has now been extended to mechanical property mapping of nanometer-resolved live cells.

Novel insights into cell mechanics can be obtained by combining these two sophisticated live cell imaging methods into a single platform, along with the capability to obtain concurrent nanometer-mechanical, biochemical, and optical data. This article demonstrates the combination, integrated performance, and superimposed data of a cellular model system using Bruker’s BioScope Catalyst™ AFM and PicoQuant’s MicroTime 200, on the basis of an IX71/81 inverted optical microscope equipped with PiFoc focus control.

Benefits of Combined Techniques

Both AFM and confocal fluorescence microscopy can be used to investigate the surface characteristics on a single molecular level. Using these techniques various aspects of the sample can be studied, for instance, surface topography can be characterized by AFM and molecular dynamics can be examined with time-resolved fluorescence spectroscopy. Combining both methods into a single system not only improves the methodology, but also creates the potential for novel investigation concepts, which are discussed in the following sections:

Live Cell Imaging

Growth and differentiation of cells are correlated with variations in cell content and shape. Synchronized acquisition of fluorescence and AFM images help in tracking changes occurring in cellular surface properties, like concentration and localization of proteins during cell development and membrane elasticity. Any topographical change occurring in cell shapes is based on changes in the cytoskeleton of cells. The rearrangement of proteins can be related to changes in localization, conformation, or orientation of proteins. The fluorophore behavior can be influenced by the difference in the protein structure as well as in the environment.

High-Resolution Molecular-Scale Imaging

The structure and packing of native as well as reconstituted membranes can be viewed through AFM. These types of membranes typically include a varied distribution of protein, sterol, and lipid components. The concurrent analysis of AFM combined with fluorescence methods could reveal the association of these structures with protein compartmentalization, and expose their function in the process.

Force and Mechanical Studies

Single-molecule force spectroscopy is yet another proven AFM method in which pairs of molecules or single molecules are interrogated mechanically to calculate intra- and inter-molecular forces. Intra- or inter-molecular distances can be measured through parallel observation of fluorescence parameters, like FRET processes caused by the movement.

Combined System Description

PicoQuant’s MicroTime 200 is an advanced confocal time resolved fluorescence microscope that comes with single-molecule sensitivity. The instrument is designed around an inverted microscope body and employs the technique of time-correlated single photon counting (TCSPC) for time-resolved data acquisition, together with picosecond pulsed diode lasers and single photon sensitive detectors for excitation.

To make image measurements, like fluorescence lifetime imaging (FLIM), a 3D piezo scanning system with a scan range of 80 x 80 x 80 µm, is integrated into the device. In the case of a typical configuration, the sample is mounted on the scanner and shifted over the immobile excitation beam. This is known as sample scanning. In the case of an alternative configuration, the objective is placed on the scanner and moved under the stationary sample. This is referred to as objective scanning.

With this unique configuration, the sample can be easily accessed from above, making it suitable for applications where unique sample compartments are required, as well as for integration with other methods requiring cryostats, patch-clamps, or AFM, as shown here (Figure 1).

Figure 1. Combined setup of the BioScope Catalyst and the MicroTime 200. The AFM with its sample stage is mounted on the inverted microscope body of the MicroTime 200, which is configured for objective scanning.

Over the last two decades, there have been continuous developments in the use of AFM for imaging living cells in near-physiological conditions. Bruker’s BioScope Catalyst™ is a high-performance AFM that has been developed specifically for biological applications. It features a flat sample scanner design that separates the piezos from any potential damage from fluids. The AFM head has an open design that provides almost unlimited physical and optical access from above the sample. The scanning range is 150 x 150 µm in closed-loop operation, with noise-free sensors to ensure accurate force measurements and excellent resolution imaging.

The tip can be moved separately with the highest range of over 20 µm (in Z-direction), allowing measurements of large-sized samples, including living cells. The AFM also comprises of a sample stage with tip-optics alignment and tip-sample alignment, making it suitable for Olympus’ IX71/81 inverted microscope body. Additionally, the AFM control software has been optimized for ease of use, including the unique ScanAsyst® AFM imaging mode from Bruker that automatically optimizes the imaging parameters for AFM.

The software also consists of microscope image registration and overlay (MIRO®) software, allowing direct correlation and calibration of AFM images, in addition to light microscopy images. Using the novel PeakForce QNM® mode, users can easily access nanomechanical data, like tip-sample adhesion, sample modulus, and dissipation by determining and examining force curves at individual image points, while imaging high-resolution sample topography.

The BioScope Catalyst AFM can be integrated with PicoQuant’s MicroTime 200 as it meets all the required needs:

  • The BioScope Catalyst is a sample-scanning AFM.
  • A scan synchronization signal is provided the controller of the BioScope Catalyst.
  • The BioScope Catalyst’s scan head mechanically fits onto the Olympus IX71/81 microscope body without affecting the operation, of normal light microscopy.
  • Adjustment of the AFM tip in x, y and z in relation to the confocal beam can be done with the sample baseplate of the BioScope Catalyst.

One of the main objectives of the combined use of the MicroTime 200 and the BioScope Catalyst is to facilitate concurrent recording of optical and AFM images of the same region of a sample. A setup should be designed where the confocal volume of PicoQuant’s MicroTime 200 can be accurately arranged with the AFM tip. One way to acquire this is to use the MicroTime 200’s objective scanning mode to accurately find the AFM tip below the optical scan range.

It is possible to move the sample baseplate and therefore, the entire AFM in x and y directions, which is handy for the pre-alignment of the cantilever tip with respect to the MicroTime 200’s optical axis. This is followed by placing the sample onto the AFM’s scan stage (Figure 2). As the AFM software directly controls all the scanning processes, the AFM’s functionality is not affected in any way.

Figure 2. Schematic of the combined setup: The sample is placed on the scanner of the AFM, and the data acquisition of the MicroTime 200 is synchronized with the scanner movement by including corresponding electronic marker signals into the collected photon data stream.

Synchronization of the Two Systems

The integration of the BioScope Catalyst and the MicroTime 200 enables simultaneous acquisition of AFM images and fluorescence lifetime. The PicoQuant TCSPC unit has a unique time-tagged time resolved (TTTR) mode that is integral to this process and helps to insert external synchronization signals or markers in the steadily recorded data stream of photon arrival times.

In the AFM scanning systems, these markers serve as signals at the start and end of individual scanning lines, which split the obtained photon stream within the scan lines and also make it possible to assign photons into image pixels. The synchronization signals, “line start” and “line stop”, can be easily accessed from the AFM controller.

Optical Alignment

Prior to beginning the measurements, care should be taken that the laser focus of the microscope and the AFM cantilever tip are properly aligned. The cantilever can be viewed via the MicroTime 200’s eyepieces. In a single eyepiece, a cross hair can be seen, and this is aligned to the microscope’s optical axis (Figure 3A). By simply moving the AFM sample baseplate, the initial coarse alignment can be done until the cantilever tip is directly placed under the cross hair (Figure 3B).

Figure 3. Cantilever with tip in the eyepiece of MicroTime 200 aligned to the cross hair (left image). The coarse positioning can be done by using the three micrometer screws at the sample stage (red arrows in right image).

Utilizing the MicroTime 200, the laser beam can be properly aligned with the cantilever tip using the acquisition of reflected and scattered light. The initial coarse alignment is enough to guide the cantilever tip within the MicroTime 200’s scan area. A superior start position in the z direction is 80 µm, which happens to be the lowest available position for the MicroTime 200 PIFOC objective nano-positioner.

In order to prevent damage to the AFM probe, small steps in Z should be carried out (about 2 µm). Figure 4 shows the resulting image. Towards the end, a bright spot in the center of the image, representing the cantilever tip, must be seen (Figure 4F).

Figure 4. Backscatter images of AFM cantilever with a SiN tip. The bright spot in the image center is the tip apex. After iteratively zooming in and scanning the AFM tip, the laser focus of the MicroTime 200 can be directly positioned at the tip apex. The sample stage is moved 2µm in z-direction for every image B-F.

Point measurements are carried out on the cantilever tip to align the laser focus with the AFM. The focal plane should be finely aligned, using the probe’s photon count rate as an indicator. It has been shown that this alignment process works with bare silicon as well as with silicon nitride AFM tips on glass bottom culture dishes and other similar glass surfaces.

In situations where the tip apex fails to provide a clear bright spot in the optical picture, the tip’s shape can be used to establish the tip apex position (Figure 5). By applying the photon count rate as an indicator, the precise confocal plane can be found again. Given that the laser power required for this calibration process is relatively low, photobleaching during the alignment procedure is not an issue.

Figure 5. Backscatter image of a probe tip with marked tip apex (red cross), which is determined by the shape of the tip base.

Imaging of Fluorescent Beads

In a primary experiment, the integrated setup of the BioScope Catalyst and the MicroTime 200 was used to image fluorescent beads (TetraSpeck, 100 nm diameter, Molecular Probes) on a glass cover slide. The results of which are illustrated in Figure 6.

Figure 6. Synchronized acquisition of fluorescence beads measuring fluorescence lifetime and topography. (A) intensity modulated FLIM image (B) AFM image and (C) merged AFM and FLIM image. Not all beads that are visible in the AFM picture also show fluorescence in the FLIM image. Scale bar is 5µm.

Using the AFM in ScanAsyst mode, the excitation wavelength was at 470 nm with an LP510 filter placed before a photon avalanche diode (SPAD) from MPD (Olympus 60X 1.2 NA water objective). In order to obtain a synchronized image, picoquant’s MicroTime 200 was utilized as a slave of the BioScope Catalyst scanner. Following a scan, an intensity modulated FLIM image can be seen in the MicroTime 200 software, including up to eight data channels in the AFM software.

The images distinctly show how both methods make it possible to image the beads, and they also reveal the variations. However, all features cannot be seen in the FLIM or AFM image (Figure 6A-C). This difference can be seen only with a synchronized acquisition method. Figure 7 shows a 3D representation of the fluorescence and AFM height image.

Figure 7. Overlay of the AFM height image as 3D representation and with fluorescence intensity in green (Image resolution 10 µm x 10 µm x 300 nm).

In order to examine the synchronization quality, a set of eight integrated AFM-fluorescence images of approximately 300 nm high fluorescent bead were obtained. The SPAD’s fluorescence channel was introduced into the AFM controller’s auxiliary photon counting input so that the intensity of fluorescence was concurrently obtained with the AFM data using the AFM software.

Using the NanoScope XY drift analysis aspect, drift rates for the AFM as well as the correlation between optical and AFM images were determined. 2.8 nm/min was the highest drift rate seen for the AFM, and 11.5 nm/min was the maximum drift rate seen for AFM-optical alignment. This indicates that the alignment between a diffraction-limited spot at high NA (for instance, 300 nm) and the AFM tip can be maintained for at least for an hour.

Fixed Cells

As the BioScope Catalyst has been optimized for biological samples, fixed glioblastoma cells or brain cancer cells were also determined. These cells were stored in PBS buffer of pH 7.2. A free diffusible GFP was expressed by the transfected cells that should be seen both in the cytoplasm and the nucleus. Using a DNA intercalating dye called DAPI, the DNA was counter-stained. The laser line utilized here does not activate the dye. The integrated setup of the BioScope Catalyst and the MicroTime 200 helped in analyzing the integrated cell topography and fluorescence lifetime of the GFP (Figure 8).

Figure 8. Synchronized acquisition of fixed glioblastoma cells expressing GFP using the MicroTime 200 in combination with the BioScope Catalyst. (A) Fluorescence lifetime distribution in the cell. (Image size 100 x 100 µm) acquired by by MicroTime 200, and aligned using MIRO within the NanoScope software. (B-H) Data taken by the BioScope Catalyst AFM in PeakForce QNM mode showing topography, and spatially resolved quantitative mechanical cell properties. (E) Photon count from the MicroTime 200, synchronously recorded by NanoScope AFM controller.

A pulsed laser at 470 nm was integrated in the MicroTime 200 for GFP excitation, and a bandpass filter HC520/35 placed before the SPAD detector was used to detect the fluorescence. The data acquired by the BioScope Catalyst in PeakForce QNM mode is shown in Figure 8B-H. If a force curve at individual image points is analyzed and measured, then modulus, topography, deformation adhesion, and dissipation of the cell can be obtained. Quantitative measurement was made through previous spring constant calibration of the AFM cantilever by utilizing the combined thermal-noise calibration technique.

Fluorescence intensity or photon count was pixel-synchronously recorded with the AFM measurement (Figure 8E) and this was done by scattering the digital APD signal of the MicroTime 200 into the pulse-counting input of the NanoScope® V AFM controller. Figure 8A displays the intensity modulated GFP lifetime dispersed in the cell obtained by the MicroTime 200. Image overlay can be done by exporting the FLIM image and superimposing it in the MIRO software from Bruker.

Both methods and instruments, the BioScope Catalyst AFM and the MicroTime 200 time-resolved fluorescence microscope, are regularly utilized for live cell studies. As a result, this evidence of performance experiment can also be applied for integrated FLIM-AFM live cell imaging.

For prolonged studies of sensitive eukaryotic cells, integrated AFM-FLIM studies can be carried out in sterilized, CO2-buffered media at physiological temperatures without introducing any environmental stress. This can be done by using the Catalyst Perfusing Stage Incubator and Catalyst Heater Stage accessories.

Conclusion

Single-molecule dynamics can be studied through the integration of time-resolved confocal microscopy and AFM. The MicroTime 200 combined with the BioScope Catalyst has been shown to be simple, easy and effective, and also eliminates the need to alter the systems. With the development of an alignment process, initial results show the capabilities of such an integrated setup. Potential applications of the AFM-FLIM technique include:

  • Live cell imaging and the analysis of the effect of protein changes in cell structure and shape and vice versa
  • Mechanical and force studies investigating intra- and inter-molecular distances using nanomanipulation and atomic force spectroscopy on the single-molecule level
  • High-resolution molecular-scale imaging by combining optically encoded functionality with sub-nanometer AFM topographic imaging

This information has been sourced, reviewed and adapted from materials provided by Bruker Nano Surfaces.

For more information on this source, please visit Bruker Nano Surfaces.

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