1.

Introduction

As noted in the 1998 National Institutes of Health Bioengineering Consortium Report: “New developments in microscopies are providing crucial information and essential approaches for understanding the structure function of cells and molecules. Molecular and cellular bioengineering is a rapidly evolving multidisciplinary area capitalizing on these technologies to create advances in research in many vital areas.” We are presenting here a state of the art technology that offers novel applications in real-time assessment of mechanotransduction dynamics in live cells by combining the structural and functional resolving power of the atomic force microscope (AFM) with rapid optical imaging at high spatial and temporal resolution provided by total internal reflection fluorescence (TIRF) and fast-spinning disk (FSD) confocal imaging. This unique integrated microscopy technology facilitates 4-D real-time biophysical studies directed toward the alteration of the intracellular force balance within cells, along with resulting functional modulations at distant sites (i.e., junctional and focal adhesions). Real-time analysis is essential for direct observations of cellular responses to mechanical or chemical stimuli, and to avoid distortion or loss of cellular components during specimen preparation, which can occur using conventional fixation and antibody-based localization studies.

The present work outlines significant improvement in our previous NanoFluor integrated microscope design¹ that combined AFM with TIRF, interference reflection microscopy, and Forster resonance energy transfer. The improved microscope is particularly well suited to study focal adhesions and cytoskeletal dynamics under AFM mechanical stimulation. By combining the AFM² with rapid optical imaging using FSD confocal imaging³ in a single instrument, substantial progress in real-time monitoring of mechanotransduction dynamics in live cells is possible. The fast-spinning disk confocal is a multibeam scanning method that permits fast scanning at high frame rates as the result of a dual-disk system, pinhole and microlens, that is synchronized with a charge-coupled device (CCD) detector. This tandem arrangement allows for enhanced detection of both low fluorescence signals and real-time image collection. In this way one can rapidly section living cells in 3-D spatial imaging over time to study the cytoskeletal dynamics with minimal exposure of cells to light, preventing cell phototoxicity. In addition, TIRF⁴ provides high contrast images that contain high resolution details of cells grown on coverslips with excellent z-resolution ( $\sim 100\mathrm{nm}$ depth). Due to minimal exposure of cells to light, TIRF images have very low background fluorescence.

Therefore, by combing these three different techniques, it is now possible to: image cellular processes with high spatial resolution or stimulate the apical cell surface using AFM; perform quantitative time-lapse imaging of the cell-coverslip interface using TIRF; and carry out rapid confocal sectioning in living cells over extended periods of time while studying molecular dynamics and protein translocation between different subcellular structures. Consequently, the integrated microscope described here offers a comprehensive view of mechanotransduction processes from the micrometer to nanometer scale in living cells.

Transmission of force is a classical physical concept applied to live cell dynamics, intracellular protein translocation, and transduction of mechanical signals along intracellular pathways. Multiple dynamic processes trigger changes in cell-cell and cell-matrix adhesions in the vascular wall, resulting in cell growth, motility, contraction, or mechanotransduction. Endothelial and vascular smooth muscle cells reside in a mechanically active environment that changes continuously. Adaptations to these local forces results in realignment of the cytoskeleton and redistribution of the numerous constituents of focal adhesions (FA) and junctional adhesion (JA) sites. The mechanism by which cells sense and respond to mechanical stimuli and the signaling pathways that drive these responses are at a relative early stage of investigation. The current understanding of the molecular processes that induce FA and JA remodeling involving force generation at a scale of pico- to nano-Newtons and a time scale of milliseconds is far from complete. The integrated instrument described here enables quantitative monitoring of rapid molecular changes and provides an important new tool for the study of transduction mechanisms initiated by agonist-induced signaling or external mechanical forces on cells. Generated data will provide new knowledge relating changes in intracellular force balance to the molecular mechanisms by which cytoskeleton communicates a chemical or mechanical stimulus to distant sites. The applications presented here employing microscopy technology that integrates optical and nonoptical scanning techniques are focused on the study of vascular cells. However, the instrument has broad applications to analysis of cellular physiology across a wide variety of adherent cell types.

The cytoskeleton and focal adhesions are dynamic structures that assemble, disperse, and turn over as cells migrate or respond to mechanical forces. They have significant roles in regulating the contractile function, cell growth, survival, and gene expression. Knowledge of the specific distribution and activity of protein constituents and their molecular dynamics will enhance our understanding of how cells detect and respond to structural and functional properties of the extracellular matrix and to neighboring cells.

The cytoskeletal network confers tensile strength to the cell. Actin, the primary cytoskeletal structural protein, together with microtubules determine cell shape. Cell shape stability depends on a network of self-equilibrating structural components, some of which are in tension and others that are in compression.⁵ Thus, in cultured vascular smooth muscle cells (VSMC) and endothelial cells, as in any other anchorage-dependent cells, external mechanical stresses are imposed on a preexisting force equilibrium generated by cytoskeletal tension.^{6, 7, 8, 9, 10, 11, 12} Forces are transmitted as solid mechanical stresses that are distributed throughout the cell via the cytoskeleton. Intracellular stresses may then be transmitted to neighboring cells via JA, and to the underlying matrix via FA at the basal cell surface.

Focal adhesions are critical points for the regulation of actin organization as well as the bidirectional transfer of mechanochemical perturbation^{13, 14, 15, 16, 17, 18, 19, 20} between the extracellular matrix (ECM) and cytoskeleton. They contain clusters of integrins (transmembrane proteins) that physically link the ECM to the cytoskeleton via a number of specific proteins, including talin, focal adhesion kinase, vinculin, $\alpha$ -actinin, and paxillin. During cellular contraction, specific proteins undergo orientation changes, become phosphorylated,^{21, 22, 23} or translocate away from the FA (e.g., zyxin^{24, 25, 26}). Which proteins are absolutely required for the formation of FA is still under investigation, and the order in which these proteins bind integrins to actin or to each other is only partially understood.^{18, 19}

At the present time it is unclear how a change in the intracellular force balance induces molecular mechanisms by which the cytoskeleton influences FA and JA reorganization and how FA and JA proteins cooperate to respond to mechanical force.

2.

Imaging across Biological Spatial and Temporal Scales

“Many of the most important mechanistic questions in cell biology might be rapidly answered if only we had molecular video electron microscopes—that is, the ability to non-destructively image specific molecules with the resolution of electron microscopy in real time.”²⁷

Important signaling events at cellular level take place in a domain smaller than $500\mathrm{nm}$ . In epifluorescence microscopy, the diffraction limits the resolution to about $250\mathrm{nm}$ in the $xy$ plane and is even lower along the $z$ axis. As mentioned before, each imaging technique has its own spatial and temporal limitations. Combinations of optical imaging techniques with atomic force microscopy may overcome some of these spatial and temporal challenges.^{28, 29, 30} Advantages of FSD confocal and TIRF imaging techniques combined with AFM are discussed next.

Fast-spinning disk confocal is a multibeam scanning method that allows rapid scanning of samples at high frame rates,^{3, 31, 32, 33} being particularly well suited to live cell imaging due to the lower levels of photobleaching and phototoxicity capabilities of this technology. This instrument involves a fully motorized Yokogawa scanning head with a dual-disk system, pinhole and microlens disks, which are perfectly aligned. The size of the pinholes is fixed, being optimal for a specific objective $(100\times 1.4\mathrm{NA})$ , and will be suboptimal for z-sectioning with a different objective. In this type of confocal microscope, the NA and the objective lens magnification are important, because the zoom-in option is not available. When compared with a classical laser scanning confocal microscope, z-resolution is limited, being dictated primarily by the spacing of the pinholes on the disk. The tandem pinhole-microlens disk partially compensate for this by directing more light through the pinholes, with the same rejection of the out-of-focus light by the disk. The actual image capture speed of this confocal system is limited only by the CCD camera used, and is discussed further in the CCD camera section.

TIRF microscopy represents a method of exciting and visualizing fluorophores present in the near-membrane region of live cells grown on glass coverslips.^{4, 34, 35, 36, 37, 38, 39} Any movement of fluorescent-tagged proteins within FA or in the immediate proximity of the cell membrane can be monitored in real time. This type of excitation can be used to obtain high contrast fluorescence images, with very low background and virtually no out-of-focus light. The confinement of fluorescence excitation to the near-field interface produces minimal exposure of cells to light in any other planes within the sample. While resolution in optical microscopy is usually diffraction limited by the wavelength of light, this optical imaging technique allows imaging of cellular structures at the coverslip-cell interface with a higher z-resolution (i.e., $<100\mathrm{nm}$ ) than confocal microscopy $(\sim 300\mathrm{nm})$ in a wide-field approach. Table 1 presents the main differences between TIRF and confocal imaging.

Table 1

Comparison of FSD confocal and TIRF microscopy. For both imaging methods, the acquisition speed is limited only by CCD camera speed and photon collection efficiency (e.g., signal-to-noise ratio).

AFM² is an attractive tool for biological applications such as live cell imaging and single molecule force spectroscopy,^{40, 41, 42, 43, 44, 45} by providing structural, mechanical and functional information at nanometer resolution under physiological conditions. The principle of operation of the AFM is very similar to that of a stylus profilometer—a sharp cantilever tip interacts with the sample surface sensing the local forces between the molecules at the tip and at the sample surface. Due to the fact that no light is involved in acquiring the sample properties, the resolution achieved by the AFM is limited only by the tip radius and the spring constant of the cantilever. When used for imaging live cells, AFM is relatively slow compared with optical imaging (minutes/image). However, when used as a force measurement tool or mechanical stimulation tool, the simultaneous use of AFM with optical imaging enables functional assessment of dynamic processes within live cells.

3.

Challenges of Integrating Fast-Spinning Disk Confocal and Total Internal Reflection Fluorescence Microscopy with Atomic Force Microscopy

Specific challenging design requirements are involved in the instrument integration process, requiring customized mechanical and optical components. Integration of the optical systems with the AFM is possible by mounting the AFM scanner on an inverted microscope platform Olympus IX-81. The system layout is presented in Fig. 1 . The key element for preserving all features of a fluorescence microscope is a customized AFM stage¹ that replaces the regular microscope stage, and an open AFM scan head design (Bioscope SZ Hybrid Scanner, Veeco Instruments, Santa Barbara, California). This type of stage is not available in any commercial AFM at this time. As configured, our microscope system allows full turret rotation, full access to the objective lens correction collars, and oil/water loading at the objective lens without moving the sample that is fixed by a magnetic holder.

Fig. 1

(a) Layout of the integrated microscope. (b) Layout of the laser-to-fiber optical train (see Fig. 2).

Fig. 2

(a) Diagram of the integrated microscope layout. The entire optical train on the FSD confocal sits on a silicon damper pad that isolates the Yokogawa unit from the optical table. Further vibration isolation is achieved by decoupling the flange between the microscope and FSD confocal, so that no direct mechanical connection exists between the two components. The microscope itself sits on adjustable legs (along the $z$ axis). The optical alignment is performed such that the primary FSD confocal camera is in register with the microscope focal plane in an open beam coupling configuration. The ZDC module is placed between the turret and the microscope objective. On the right-hand side, we placed the TIRF illuminator and its CCD camera. (b) Diagram of the laser-to-fiber optical train. FS is field stop, and POL-polarizer. The optical filters are listed in Table 2 and the main components are listed in Table 3.

Table 2

Optical filters. The far right column of numbers relates to the part numbers in Fig. 2. All fluorescence filters were made by Chroma Technology (Brattleboro, Vermont).

Table 3

Component list of the integrated microscope. The far right column of numbers relates to the part numbers in Figure 2.

Special attention was paid toward insulating the integrated microscope from building and acoustic vibrations that can affect or even destroy the AFM measurements. The integrated microscope and the laser head are placed on a research-grade vibration-insulation tabletop (Newport, Irvine, California). A detailed presentation of vibration isolation, cooling, and electrical wiring requirements were presented elsewhere.¹ While camera cooling fans and other drivers containing fans and moving parts were isolated from the main vibration-isolation table by placing them on a self-standing shelf, this is not possible for the Yokogawa scanner head. The scanner that rotates at $5000\mathrm{rot}∕\min$ must be placed in the immediate vicinity of the microscope, having its optical path aligned with the microscope optical path. This important source of vibration was eliminated by designing an open-coupling flange between the scanner head and the microscope, and by placing the scanner on a silicon damper pad that isolates the scanner from the tabletop [Fig. 2a ].

The optical lever principle used in the AFM scanner to detect the cantilever deflection consists of a laser beam coming from a laser diode in the scanning head that is reflected off the back of the cantilever onto a photodetector. The laser diode emits at $693\mathrm{nm}$ and was chosen far from the spectral range of wavelengths used for fluorescence measurements. It is known that laser diodes have a broad emission spectrum. To optimize the spectrum output and minimize its interference with the fluorescence, a notch optical filter was mounted into the AFM scanner (Fig. 3 ). Moreover, even filtered in this way, light coming from the laser diode reaches both cameras, quenching the dim fluorescence, and also reaches the oculars, making impossible the use of microscope eyepieces while the AFM is turned on. Thus, a different filter was designed to block the light of the laser diode and was placed in front of both cameras and oculars. A limitation introduced by this filter results in blocking the visible spectrum in the red wavelengths $(>670\mathrm{nm})$ , such that dyes like CY5 cannot be used in simultaneous measurements with AFM. This limitation is easily overcome when AFM is not in use, by placing these blocking filters in specially designed sliders that allow sliding the blocking filter in and out of the optical path for TIRF and FSD confocal. For alignment purposes of the AFM optical lever, it is useful to see some of the laser diode light on the video camera used for the optical lever alignment. This required the design of a different optical filter that partially blocks the diode beam in front of the video camera, such that only a small spot of light is visible in the field of view, allowing for a precise alignment of the optical lever.

Fig. 3

(a) Transmission curves for the optical filters necessary to exclude the AFM laser diode from the fluorescence path: solid black line—ET670sp is placed in front of the EMCCD and eye pieces; gray line—ET680sp is placed in front of the video camera; dotted line—notch filter in the AFM head; dashed line—generic spectra of a laser diode emitting at $690\mathrm{nm}$ . The laser diode spectrum is wide enough to bleed through into the visible spectrum suppressing the fluorescence signal. (b) The AFM laser diode light captured by the EMCCD without the notch filter in place (to be able to capture an image that does not completely blank the EMCCD, an extra $647\text{-}\mathrm{nm}$ filter was set in the optical path). (c) When the notch filter is mounted in the AFM head, the laser diode light is suppressed. Both images in (b) and (c) were captured with the protection filter in place (solid black line in (a)). (d) The laser diode light is visible to the video camera for an easy alignment of the optical lever [video filter in place is the gray line in (a)].

A user-friendly design of the filter combinations necessary for different applications with the spinning disk confocal required an external filter wheel (Sutter Instruments, Novato, California) to be mounted at the out-put path of the scanning unit. Therefore, different combinations of two or three channels for confocal imaging are possible without moving the filters in the optical path while scanning. All optical filters were provided by Chroma Technologies (Brattleboro, Vermont) and are included in Table 2 . The extension of the optical path to the camera was possible by using an optical extender from Optical Insights (Tucson, Arizona) that ensured a good focused image projected on the camera chip without introducing optical aberrations.

A z-drift compensator (ZDC) mounted between the turret and objective nosepiece [Fig. 2a] allows for locating the specimen-coverslip interface with high precision. The ZDC module uses a $785\text{-}\mathrm{nm}$ laser diode to find with high precision the upper surface of the coverslip. The ZDC autofocus routine is faster than a conventional microscope autofocus, and is not affected by changes in contrast or fluorescence intensity in the filed of view. Thus, ZDC represents an important addition, providing an optimal solution for long-term time-lapse TIRF experiments where maintenance of the focal plane is critical. Note that TIRF laser alignment is affected by the ZDC module. If the ZDC is to be employed, the laser alignment must be performed with the ZDC module in the optical path. The ZDC autofocus is also used in time-lapse confocal imaging.

4.

Description of the Integrated Microscope System

The overall schematic of the integrated microscope shown in Fig. 1 is presented in Fig. 2, and a detailed list of components is presented in Tables 2, 3 . Additional information regarding laser illumination and image acquisition systems are presented next.

5.

Single-Cell Applications

5.2.

Live Cell Imaging

In this section, several applications of the integrated AFM-optical imaging microscope are presented.

In Fig. 4 vascular smooth muscle cells (VSMC) were imaged in AFM mode to show cell topography. TIRF and confocal images of the same cell, transfected as above, were acquired immediately following acquisition of the AFM image. An excellent registration between the AFM image and the TIRF and FSD confocal fluorescence images can be achieved as shown in the overlays [Figs 4d and 4e]. AFM provides a general topographic view of the cell with few actin cytoskeletal fibers immediately beneath the membrane. The confocal image reveals a multitude of actin fibers aligned along the cell axis. If TIRF images are compared with FSD confocal images of the same cell, more background from the cell body is shown by the confocal images. Not all actin fibers are present in the AFM image, because the AFM tip senses only features immediately beneath the cell membrane. The fluorescence from the actin fibers (actin-mRFP) inside the cell body, as well as focal adhesions (vinculin-GFP) are obvious in the confocal and TIRF images. TIRF images were acquired sequentially with an exposure time of $100\mathrm{ms}$ , and FSD confocal images were acquired as a two-channel 3-D stack of 20 planes with the same exposure time and presented as maximum projection.

Fig. 4

Same VSMC transfected with mRFP-actin and vinculin-GFP was imaged by: (a) AFM at the apical cell surface; (b) TIRF at the basal cell surface; and (c) FSD confocal throughout the cell body is shown as maximum projection of a 3-D stack. Excellent overlaps between the AFM and TIRF or FSD confocal images are shown in (d) and (e), respectively. The stress fibers present in the AFM image overlay with actin fibers from the fluorescence images. Image size is $66\times 64\mu \mathrm{m}$ .

Figure 5 shows confocal images of VSMC subjected to mechanical stimulation applied by AFM to the apical cell surface. The mechanical stimulation was induced by a functionalized AFM probe ( $2\text{-}\mu \mathrm{m}$ glass bead coated with fibronectin), which was set at a chosen $xy$ coordinate on the cell surface and moved upward in discrete steps as described in Ref. 1. Following each controlled upward movement of the functionalized cantilever tip along the $z$ axis (the position of which is shown by white dashed lines), the cell responds to the mechanical stimulation. Confocal images of the same VSMC, transfected with actin-mRFP, before and after AFM stimulation, show that the cell significantly modified its shape, such that actin filaments reorganized in almost a regular pattern of parallel actin bundles following the long axis of the cell. Moreover, the $xz$ and $yz$ side view images of the same cell (see arrows) obtained by 3-D image reconstruction from consecutive confocal sectioning of the cell show an increase in cell height after the mechanical stimulation, as well as a redistribution of the actin filaments toward the bottom of the cell.

Fig. 5

VSMC transfected with mRFP-actin were imaged by FSD confocal before (a) and after (b) AFM mechanical stimulation. Following mechanical stimulation, actin filaments reorganized in almost a regular pattern of parallel bundles along the cell axis. The side views show actin reorganization and recruitment toward the cell bottom along the indicated section in the $xz$ and $yz$ planes (see arrows). White dashed lines represent the AFM tip above the cell. Image size is $50\times 86\mu \mathrm{m}$ .

Videos 1 shows real-time cell shrinkage and FA rearrangement due to AFM mechanical stimulation. TIRF images of VSMC transfected with actin-mRFP and vinculin-GFP show that F-actin is recruited at the bottom of the cell in the immediate vicinity of the FA that also reorganizes to better anchor the cell to the substrate to oppose the applied force. The TIRF images were acquired during the AFM stimulation experiment with $100\text{-}\mathrm{ms}$ exposure time at eight time points between 0 and $80\min$ . The video is shown at $1.5\text{frames}∕\sec$ .

Video 1

Real-time cell restructuring shown as time-lapse images acquired in TIRF mode for the duration of the experiment $(\sim 80\min )$ . VSMC were transfected with vinculin-GFP and actin-mRFP. The images were acquired with $100\text{-}\mathrm{ms}$ exposure time at eight time points between time 0 and $80\min$ . The video is shown at $1.5\text{frames}∕\sec$ . Image size is $91\times 91\mu \mathrm{m}$ . (QuickTime, 2.36 Mb).

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Videos 2 shows real-time actin restructuring due to AFM mechanical stimulation. Time-lapse 3-D stacks were acquired using FSD confocal for the duration of the experiment $(\sim 80\min )$ . The arrows in Fig. 6 (the first frame of the video) indicate actin fibers that disappear (a), thin out due to actin restructuring (b), and bend due to the mechanical stimulation (c). The AFM tip above the cell is shown by white dashed lines. Images were acquired with $100\text{-}\mathrm{ms}$ exposure time as a 3-D stack of 20 planes at six time points between time 0 and $80\min$ . The video is shown at $1\text{frame}∕\sec$ .

Fig. 6

A VSMC transfected with mRFP-actin was imaged by FSD confocal. The arrows indicate actin fibers that show remarkable restructuring: (a) actin fibers disappear, (b) actin fibers thin out due to actin restructuring, and (c) actin fiber bends due to the mechanical stimulation (see text). The AFM tip above the cell is shown by white dashed lines. Image size is $91\times 91\mu \mathrm{m}$ (see Videos 2)

Video 2

Real-time actin fibers restructuring shown as 3-D time-lapse video from FSD confocal imaging of VSMC transfected with actin-mRFP. The images were acquired with $100\text{-}\mathrm{ms}$ exposure time as a 3-D stack of 20 planes at six time points between time 0 and $80\min$ . The video is shown at $1\text{frame}∕\sec$ . A small reflection of the fluorescence due to the AFM tip is also visible. (QuickTime, 1.75 Mb).

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The basal cell area was reduced $8.7\pm 2.1\%$ after AFM mechanical stimulation, but the vinculin area increased $11.1\pm 2.1\%$ $(\mathrm{n}=5)$ . Also, F-actin at the basal cell area (TIRF images) increased $16.9\pm 1.9\%$ and F-actin in whole cell volume (confocal images) increased $8.2\pm 0.3\%$ $(\mathrm{n}=4)$ . By recruiting FA proteins and F-actin at the basal cell area, the cell reinforces its attachment to the substrate to better resist the mechanical stimulation. Area measurements were performed in SlideBook software (Intelligent Imaging Innovations, Denver, Colorado). The percent change in protein area was calculated as percent $\text{change}=[\left(\text{treatment}\text{-}\text{control}\right)∕\text{control}]\times 100$ . Therefore, a significant rearrangement of the actin filaments and FA took place due to the local mechanical cell stimulation at the apical cell membrane that induced changes into the cellular structure throughout the cell body.

Examples of the utility of the integrated microscope to examine the restructuring of actin filaments and FA as the result of mechanical stimulation to the surface of cells have been presented. Another application under development is the analysis of FA reorganization as a function of forces generated from within cells. Videos 3 and Videos 4 show calcium oscillations in liver epithelial cells Clone 9 (ATCC, Manassas, VA), the frequency of which is dependent on agonist concentration as previously characterized.⁴⁸ By treating the cells with $2\text{-}\mathrm{nM}$ vasopressin, intracellular calcium oscillations are induced at about $50\mathrm{mHz}$ . Videos 3 was acquired as time-lapse 3-D stack for each time point at $3\text{frames}∕\sec$ in FSD confocal microscopy. For Videos 4, we were able to acquire images as fast as $50\text{-}\mathrm{ms}$ exposure time with no binning and good image quality, which translates to $20\text{frames}∕\sec$ and shows details of two calcium oscillations in the same cell. These frequency-encoded calcium signals activate key signaling pathways that mediate myosin-light-chain-dependent contractions, which are transmitted by FA to the extracellular matrix, and can be studied by AFM-optical imaging combined experiments.

Video 3

Time-lapse movie of FSD confocal images of calcium oscillations in liver epithelial cells, induced by vasopressin treatment, acquired with $60\text{-}\mathrm{ms}$ exposure time. The video is shown at $5\text{frames}∕\sec$ . Image size is $40\times 28\mu \mathrm{m}$ . (QuickTime, 1.66 Mb).

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Video 4

Time-lapse video of FSD confocal images of calcium oscillations in a single liver epithelial cell, induced by vasopressin treatment, acquired with $50\text{-}\mathrm{ms}$ exposure time at $20\text{frames}∕\sec$ . Image size is $37\times 35\mu \mathrm{m}$ . (QuickTime, 8.88 Mb).

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6.

Concluding Remarks

Cells have dynamic structures that adapt continuously to their micromechanical environment. The importance of the external mechanical stimuli that determine profound changes intracellularly can be seen in pathological conditions as hypertension, atherosclerosis, or inflammation. Fundamental understanding of the mechanotransduction process, and of the cellular pathological responses, will allow new insights into understanding the structure-function relationships of cells. The most interesting cellular behaviors are highly dynamic and occur locally in specific cell regions: cell signaling response to mechanical stimulation, FA and actin turnover, protein translocation, conformational changes of proteins, etc. Thus, particular cellular phenomena have their own time and distance scales that have to be matched by the instruments that we use to analyze them.

The technology presented here is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells. By integrating in one instrument the AFM with TIRF and FSD confocal microscopy, we are able to follow and quantify specific adhesion proteins or cytoskeletal rearrangement and cell signaling in real time. This instrument enables future studies of 3-D spatial optical imaging of molecular dynamics in different subcellular structures at a time scale of milliseconds while applying mechanical stimulation forces in the range of pico- to nano-Newtons. Cell response to various types of agonist or mechanical stimulation can be observed directly, and the dynamics and characteristics of reorganization at subcellular level may be studied in real time.