Introduction To The Mechanical Properties Of Living Cells Using Atomic Force & Fluorescence Microscopy Mark Murphy, David R Burton, Mike Lalor, Francis Lilley, Catherine Randall GERI, Liverpool John Moores University, UK
[email protected]
Francis Manning, Steven Crosby School of Biomolecular Sciences Liverpool John Moores University, UK
Abstract The atomic force microscope (AFM) has become an increasingly useful tool for the life sciences. One of the great advantages of AFM is its ability to image and measure forces in living biological samples at subnanometer resolution in a physiologically stable environment. However, imaging living cells can be extremely challenging and requires consideration of sample preparation methods and stabilisation of the AFM system. In addition, the parameters used for imaging must be empirically determined for each cell type. This talk will aim to introduce the basic concepts of AFM and highlight its ability to image living cells. The results of experiments will be presented in which two cell types with distinctively different morphologies: fibroblastic and epithelial, were examined by AFM in both contact and tapping mode. In addition, this talk will introduce fluorescence microscopy and the concept and uses of AFM force measurements on live cells in order to investigate the role of the cell’s cytoskeleton on the mechanical properties of living cells.
air and liquid. However, achieving optimal performance can still be extremely challenging even for more experienced users.
Laser Photo Detector Z-piezo Sample X-Y
Stage
Objective Lens Fig 1. Schematic diagram of AFM operation
Key words
1.1 Live Cell Imaging
Atomic Force Microscopy, Fluorescence Microscopy, Cell, Cytoskeleton, Image Parameters
Live cell imaging with AFM requires the maintenance of stable physiological conditions and a stable AFM set-up. Temperature is usually controlled by the use of selected heating methods (e.g. heated sample stages) to help keep cells viable. While physiological stable buffers or CO2 control can be used to maintain a stable pH [Le Grimellec et al., 1997, Hassan et al., 1998, Le Grimellec et al., 1998, Nagao et al., 1998, Vié et al., 2000]. System stability is achieved by isolating the AFM from external vibration and acoustic noise [Schaus, and Henderson 1997] which if not eliminated will affect image quality. This is usually achieved using anti-vibration tables and acoustic isolation enclosures.
1. Introduction Since its invention in 1986 [Binnig et al., 1986] the atomic force microscope (AFM) has become an important tool in many areas of research and the last decade in particular has witnessed a large increase in the number of publications relating to the biological sciences. This is mainly due to the AFM’s ability to image and measure forces of biological samples in a physiologically stable environment [Le Grimellec et al., 1994, Hansma et al., 1996, Le Grimellec et al., 1998, Vié et al., 2000, Mücke et al., 2004]. AFM imaging of live cells is now performed routinely in
The AFM utilises a sharp probe attached to one end of a cantilever (Fig 1) and can be operated in both contact and tapping mode (AC). In contact mode the
which there are three types; microtubules, actin filaments and intermediate filaments which collectively form the ‘cytoskeleton’ (Figs 3 & 4). The cytoskeleton spans the cytoplasm (internal viscous fluid) and interconnects the cell nucleus (DNA containing body) with the extracellular matrix. The cytoskeleton is involved in virtually all cellular processes and abnormalities in the cytoskeleton frequently result in disease such as cancer [Frans and Bosman 2004]. In normal cells the cytoskeleton is highly organised while in cancer cells the cytoskeleton becomes dynamically deregulated and highly disorganised, ultimately leading to aberrant cell migration (spread of tumours throughout the body) [Lambrechts et al., 2004].
Indentation (nm)
1
Deflection (nm)
AFM probe is kept in constant contact with the sample at a predefined set-point deflection, which is maintained by use of a z-piezo crystal and an electronic feedback system. In AC mode the cantilever is oscillated usually at or near its resonance frequency at a predefined set-point amplitude. Deviations from the set-point values during imaging are recorded and the image is digitally constructed. AFM imaging of living cells is complicated due to their dynamic viscoelastic properties, heterogeneous surfaces and the requirements for a strong adherence between the cells and their underlying substrate. Variation in morphology further complicates live cell imaging. Some cells flatten down to their substrate making them ideal for AFM imaging such cells include fibroblasts [Nagao et al., 1998, Le Grimellec et al., 1998, 1997]. Fibroblast cells are spindleshaped cells which make the structural fibers and ground substance of connective tissue. When grown on a substrate treated with adhesion promoting factors fibroblasts can be imaged quite readily. In contrast, epithelial cells provide more of a challenge. Epithelial cells cover or line all internal and external body parts and can possess a squamous, cuboidal or columnar morphology. Successful imaging of epithelial cells usually dictates that the cells be grown as a monolayer on a treated surface. This helps provide a reasonably homogeneous surface across the cells and strong adhesion between cells, which helps to withstand the actions of the scanning cantilever. Epithelial cell processes such as active secretion and protrusive membrane features also make live cell imaging difficult especially in AC mode. During oscillation, on its downward movement the tip may stick to the surface making imaging impossible. In contrast fibroblastic cells appear quite smooth and imaging these cells in AC mode can be readily achieved [Rotsch et al., 1997, Le Grimellec et al., 1997]. Because of the problems outlined above with AC mode most workers tend to image live cells using contact mode with soft cantilevers in the range of 0.01-0.38 N/m [Hoh and Schoenenberger 1994, Hassan et al., 1998, Le Grimellec et al., 1998, Rotsch et al., 1999, Pesen and Hoh 2005].
2 3 0
2
1
3
0 Z-Movement
Fig. 2 Diagram of AFM force curve taken on a hard substrate such as glass (2) and a cell (3). 1 = noncontact region
1.2 AFM Force Measurements The AFM can also be used to investigate mechanical properties of cells by use of ‘force versus distance’ curves as the cell is indented with the AFM probe (Fig. 2). From the resulting curve, indentation versus force can be plotted and this curve fitted to an appropriate model (e.g. Hertzian model of contact mechanics) to determine properties such as elastic modulus. The mechanical integrity of animal cells is governed largely by a network of internal protein polymers of
Fig 3 Fluorescence microscopy images of LL24 fibroblast cells obtained using fluorescent probes. Green = actin filaments, red = microtubules and blue = DNA in the cell nucleus. Intermediate filaments are not shown.
2.2 Fluorescent Microscopy Cells were fixed (dehydrated) using methanol and incubated for 1 hr with fluorescence antibodies one specific for actin and one specific for microtubules. For labelling DNA cells were incubated with a fluorescence dye with a binding specificity for DNA.
2.3 Atomic Force Microscopy
Fig 4 Fluorescence microscopy images of NCI H727 epithelial cells obtained using fluorescent probes. Green = actin filaments, red = microtubules and blue = DNA in the cell nucleus. Intermediate filaments are not shown. Little is known about the mechanical properties of cells and how mechanical properties may differ between different cell types and normal cells and cells of a pathological nature such as cancer cells. To date, research into the mechanical properties of cells has relied heavily on techniques such as optical tweezers. However because of its sensitivity, accuracy and ability to image and measure forces in the piconewton range the AFM is rapidly becoming the tool of choice. The work outlined below is a result of initial experiments to [1] determine empirically the optimum scan parameters needed to obtain high quality images of two types of live human cells with distinctly different morphologies. [2] Use fluorescent microscopy to visualise actin and microtubule filaments. The main scan parameters include setpoint, proportional and integral gains, scan rate (Hz) and scan size. Set-point is proportional to loading force, which is in turn related to cantilever spring constant (cantilever stiffness N/m). Proportional and integral gains determine the response time of the feedback system. With atomic force microscopy the higher the gain values the quicker the response time of the z-piezo movement.
2. Materials and Methods 2.1 Sample preparation Cells were grown on treated glass slides to promote cell adhesion under controlled conditions (37ºC, in a humidified 5% CO2/95% air atmosphere).
All images presented were obtained using a Molecular Force Probe-3D (MFP-3D) atomic force microscope (Asylum Research, Santa Barbara, Ca) with software written in IGOR pro (Wavemetrics Inc., USA). The MFP-3D is equipped with a 90 µm x-y scanning range, z-piezo range >16 µm and was coupled to an Olympus IX50 inverted optical (IO) microscope. The MFP-3D-IO was placed upon a TS150 active vibration isolation table (HWL Scientific instruments GmbH, Germany), which was located inside an acoustic isolation enclosure (IGAM mbH, Germany) to help eliminate external noise.
2.3.1 Contact Mode Imaging V-shaped, silicon nitride cantilevers (spring constant 0.02 N/m, length 200 µm, OMCL-TR400PSA-1, Olympus) were used for contact mode imaging. Before imaging the cantilever was positioned above a glass slide in Phosphate Buffered Saline (PBS) at room temperature to thermally equilibrate (30 mins). The glass slide was then replaced with the sample under PBS and left for a further 30 minutes. Once stable, the optical lever sensitivity was determined on a bare region of the glass substrate from force versus distance curves taken at 1 Hz, (1 force curve per second) with the scan size set at 0 µm (no XY tip movement). Determination of optical lever sensitivity is necessary in order to convert the voltage applied to the piezo into a deflection distance of the cantilever in nanometers. To accurately determine the cantilever spring constant the resonant frequency of the cantilever was measured using the thermal noise method which is software driven for the MFP-3D. Briefly, the cantilever is acoustically excited causing it to oscillate at its natural frequency. This coupled with the optical lever sensitivity is used to determine cantilever stiffness in N/m. In determining loading force, force curves were taken (1 Hz) on cells prior to imaging without changing the tip or the position of the laser. Force curves allow the control of loading force applied to the sample before and during imaging thus helping reduce the chances of sample damage or displacement. For imaging the fibroblasts after engaging the tip to the surface the loading force was lowered to the desired set-point voltage by decreasing the set-point
value until the quality of the images were optimal. This corresponded to loading force values between 2 - 2.8 x 10-9 N. In order to minimize sample damage due to the relatively large scan sizes (27-49 µm), scan rates were kept below 1 Hz resulting in approximate scan velocities ranging from 27-79 µm/s respectively. Proportional and integral gains were increased during imaging to the value just below where the feedback started to oscillate. For epithelial cells, loading force was lowered and maintained between 0.1- 0.7 x 10-9 N in order to achieve good images. All contact images of epithelial cells were carried out below 1 Hz with the scan size ranging from 39-83 µm which corresponds to approximate scan velocities of between 73-155 µm/s. Proportional and integral gains were adjusted as described above.
2.3.2 Tapping Mode Imaging For tapping mode (AC) imaging sample preparation was the same as described with contact mode. Vshaped silicon nitride cantilevers (spring constants 0.32 and 0.58, length 56 & 57 µm, DNP series, Vecco Instruments) were used for all images. Optical lever sensitivity was determined from amplitude calibration plots (a-d plots, 1 Hz) on a bare region of the glass substrate with the scan size set to 0. The slope of the a-d plots was used to calculate the cantilever oscillation amplitude in nanometers [Vié et al., 2000]. As with contact mode the thermal noise method was used to determine the resonant frequency and spring constant of the cantilevers. The cantilevers were tuned to a resonance centred between 100-113 kHz. Cell imaging was performed with an oscillation magnitude, which was damped to approximately 9 nanometers for LL24 cells and 39 nanometers for NCI H727 cells, corresponding to a damping ratio of 26:78 % of the free-air amplitude, respectively. Scan rates for imaging LL24 cells was adjusted to 0.4 Hz to compensate for a large scan size of 89 µm (scan velocity 78 µm/s). For epithelial cells the scan rate was maintained at 0.6 Hz with a scan size of 26 µm (scan velocity 33.61 µm/s). Integral and proportional gains were set in a similar manner as those described for contact mode.
Fig 1 Fig 2 Fig 3 Fig 4 Fig 5 Fig 6 Parameter LL24 LL24- NCI-C NCI-C LL24 NCI-
Scan Size (µm) Scan Rate (Hz) Scan speed Spring Constant Loading Force Int. Gain Prop. Gain
-C 49
C 27
83
39
-AC 89
AC 26
0.8
0.5
0.9
0.9
0.5
0.6
79
27
155
73
78
33
0.02 0.02
0.02
0.02 0.32
0.58
2 x 2.8 x 0.7 x 0.1 x 10-.9 10-.9 10-.9 10-.9 1.7 2.2 0.9 0.9
*
*
7.2
6.3
0.05
0.2
0.09
0.2
0.05
0.05
Table 1. Summary of scan parameters. The table shows the optimised image parameters, which were needed in order to produce high quality images. Optimal scan parameters were determined empirically. C = contact mode, AC = tapping mode
5 (a)
5 (b)
3. Results 2 (a)
6 (a)
Figure 5 (a) shows an AFM deflection image of a lung fibroblast cell (LL24). Figure 5 (b) shows the corresponding 3-D plot constructed 8 (athe ) AFM height data. from
6 (b)
8 (b)
5 (b)
7 (a)
9 (a)
9 (b)
7 (b)
6 (b) 4 (b)
10 (a)
10 (b)
Figure 10 (a) shows an AFM amplitude image of lung epithelial cell (NCI H727). Figure 10 (b) shows the corresponding 3-D plot constructed from the AFM height data.
3. Discussion To better understand AFM imaging of living cells in vitro, two cell types, with completely different morphologies were chosen, a fibroblastic cell line (LL24) and an epithelial cell line (NCI H727). Cells with a fibroblastic morphology are generally considered more suitable for imaging and appear quite frequently in AFM publications. In contrast, imaging live epithelial cells provides more of a challenge, which may explain the low number of published images of these cell types. For both cell types, contact mode and tapping mode imaging was carried out and the results compared.
3.1 Contact Mode AFM Figs. 5 (a) and 6 (a) show contact mode deflection images of human, primary lung fibroblast cells (LL24) and Figs 5 (b) and 6 (b) the 3-D reconstructed isometric plots. Maximum z-height of each cell was obtained from the AFM height images and was approximately 2 µm. Although the images look different, sample preparation was consistent, including seeding density and seeding time (5 x 104 cells/cm2 and 48hrs, respectively). However, scan parameters differed between images (see Table 1). Fig.5 was captured using a larger scan size than Fig. 6 (49 µm versus 27 µm). Scan rates and thus scan velocities also differed between images, 79 µm/s (0.8 Hz) for Fig. 5 compared with 27 µm/s (0.5 Hz) for Fig. 6.
It is often stated that larger scan sizes require smaller scan velocities in order to help maintain a good tracking between the tip and sample and to help reduce the risk of damaging soft samples. This was not true in this instance and is probably due to the overall scan velocities being relatively low compared to those typically used in contact mode imaging. Proportional and integral gains also differed between the two images. Fig. 5 was captured with proportional and integral gains of 0.05 and 1.7 respectively, while fig. 6 required gains of 0.2 and 2.2. It is generally thought that faster scan velocities require larger gain settings in order for the tip to track surface features accurately. This was not true in this case as fig. 5 was captured with a considerably faster scan velocity than fig. 6 but required lower gain values (Table 1). This is probably due to the heterogeneous surface visible in fig.6 as compared to fig. 5 and indicates that choice of gain values may depend on surface morphology. Fig. 6 displays more fine detail than Fig. 5, which probably reflects the differences in scan size, imaging force and scan speed. Both images were captured with 256 x 256 data points, therefore the smaller scan size used for fig. 6 has resulted in greater resolution. A cantilever with a stiffness of 0.02 N/m was used for both images. However, fig. 5 was captured using a scanning force of 2 x 10-9 N, while fig. 6 was imaged using a force of 2.8 x 10-9 N. The apparent presence of stress fibres throughout the cell may be due to the larger force used for imaging compared with fig. 5. This would fit with the ‘draping model’ proposed by Henderson et al., (1992) which states that the applied vertical force of the tip causes the membrane to “drape” over the rigid subcellular structures during imaging. It is difficult to determine if the underlying structures visible in fig. 6 are due to force alone, especially as the force difference used for the two images was not too dissimilar. With respect to scan speed, Schaus et al., (1997) found that cytoskeletal elements were not as prominent in AFM contact images when the scan velocity was increased. This is probably due to the feedback system not being able to respond quickly enough to the changes in surface topography while scanning at high speed. This may be the case here and it is possible that lowering the scan speed may have resulted in fig. 5 showing more fine detail. Bushell et al., (1999) found that fibroblast cells grown on glass coverslips lack submembranous detail with images displaying only the outline of the cell. In contrast, when grown on treated substrates to promote cell attachment images showing underlying cytoskeleton features were obtained. As both cells were grown on treated substrates in this study it is possible that the level of cell attachment was greater for fig. 6 even though the maximum z- height for both cells was almost identical. It may also be the case that the two cells
were at different stages in their cycle. This suggests that difference in the scan force, scan velocity, scan size and natural differences such as level of cell attachment or cell cycle stage may have all contributed to the presence of the stress fibres visible in fig. 6 and the absence of stress fibres in fig 5. Figs. 7 (a) and 8 (a) show contact mode deflection images of epithelial cells (NCI H727) and figs. 7 (b) and 8 (b) the corresponding 3-D reconstructed isometric plots. Both images were captured using 512 x 336 data points with the scan size differing dramatically, 83 µm versus 39 µm, respectively. Scan velocities differed greatly between images 155 µm/s (0.9 Hz) for fig. 7 compared with 73 µm/s (0.9 Hz) for fig. 8. Proportional and integral gains were identical for both images (0.05 and 0.9, respectively). Fig. 7 shows a cluster of epithelial cells, while fig. 8 shows a single epithelial cell growing on the same substrate. Maximum z-height for these images is 4.2 and 2.9 µm respectively, with the difference in height probably reflected in cell density i.e. the greater the cell density the greater the height. When growing at high density NCI H727 cells pack together and will eventually form a monolayer. This gives them a columnar morphology, thus increasing cell height. Electron microscopy studies of NCI H727 monolayers suggest an average height value of approximately 10 µm (data not shown). In contrast, when NCI H727 cells are at low density, morphology takes on a more flat appearance as the cells have room to spread. By comparing the deflection images it is seen that fig. 8 (a) shows fine, hair-like surface features giving the image a fuzzy appearance. The bottom right-hand corner of fig. 8 (a) also shows the presence of a protruding structure above a fine filament, which was observed to move backward and forward in the direction of the scan. NCI H727 cells are secretory cells [Pu et al., 1997] and hence will contain cell surface features to facilitate this. Also, like other epithelial cells NCI H727 may contain membrane features such as glycocalix [Le Grimellec et al., 1998]. None of the structures in the images have been identified, but protruding structures have been reported as ‘spikes’ in other AFM images of epithelial cells [Hoh and Schoenenberger 1994 ]. These structures are not visible in Fig. 7 (a), and with respect to the fine hair-like features may be due to the difference in resolution. As mentioned previously increasing scan speed has been shown to reduce the presence of fine detail in AFM contact mode images. Therefore it is also possible that the faster scan speed used in fig. 7 may also have contributed to the lack of fine detail in fig. 7. Both images were obtained using cantilevers with the same stiffness (0.02 N/m). However, fig. 7 was
imaged using a loading force of 0.7 x 10-9 N while fig. 8 only 0.1 x 10-9 N respectively. This suggests that the difference in the loading force used for figs. 7 and 8 may have also resulted in fig. 7 lacking fine membrane detail. When the force is too high the action of the scanning cantilever will compress and push aside soft membrane structures. This coupled with the smaller scan size and slower scan speed may be why fine surface detail or the presence of protruding ‘spikes’ are apparent in fig 8 but not fig. 7. Such features are all but lost from the reconstructed height images except for the ‘spike’. Unlike the case for the fibroblast images, fig.7 and fig. 8 did not require a difference in gain settings. This is most likely because the surface of figs. 7 and 8 are relatively homogenous as compared to fig.5 and fig. 6, which are completely different and did require different gain settings. Also, the loading forces used when imaging the epithelial cells as compared to the fibroblast cells were very small. This was due to the difference in cell morphology. When higher forces were used as with the fibroblasts, imaging proved extremely difficult even when grown in a monolayer and good quality images proved quite rare. This is largely due to the epithelial cells not being as robust as fibroblasts and not adhering as strongly to the substrate as the fibroblasts. This often resulted in cells becoming dislodged from their substrate or damaged, ultimately resulting in tip contamination.
3.2 Tapping Mode AFM Fig. 9 (a) shows a tapping mode, amplitude image of a layer of LL24 fibroblast cells. Fig. 10 (a) shows a tapping mode, amplitude image of a single NCI H727 epithelial cell. Both images have been reconstructed using 3-D isometric plots and maximum heights for both images are 2.8 µm and 2.3 µm, respectively. Figs. 9 and 10 were captured using cantilevers with spring constants of 0.32 and 0.58 N/m respectively. When softer cantilevers were used as in contact mode they were observed to stick to the cell surface on their downward movement. This was particularly noticeable when imaging epithelial cells and was evident from observing the deflection signal in the sum and deflection meter displayed in the MFP-3D software. In this study it was found that, in tapping mode, cantilevers with a spring constant below 0.32 N/m provided no images of the LL24 fibroblast cells. In contrast, cantilevers with a spring constant below 0.58 N/m provided no images of the epithelial NCI H727 cells, irrespective of scan parameter settings. Fig. 9 was imaged with an oscillation magnitude that was damped to 9 nanometers, while Fig. 10 required an oscillation magnitude damped to 39 nanometers.
Free-air amplitudes for Fig. 9 and 10 were approximately 34 and 50 nanometers, respectively. The larger free-air and cantilever damping used for imaging the epithelial cells was required to overcome ‘adhesive’ membrane properties and surface features as seen in Fig 8 (a). As mentioned previously NCI H727 are secretory cells and may be actively secreting during imaging. This coupled with surface features such as glycocalix can disrupt the mechanical resonance of the cantilever, resulting in poor quality images when operating in AC mode. Many attempts were made to image epithelial cells in tapping mode with the overall image quality generally poor. Unlike contact mode images, tapping mode images of epithelial cells resulted in images showing smooth bulges on the cell surface (Fig 10 a). It is not clear what these structures represent or if in fact they are artefact but smooth bulges have been reported in contact mode images of other epithelial cells [Hoh and Scoenenberger 1994]. In contrast, the LL24 fibroblasts are not secretory cells and the membrane surface is much smoother compared to the NCI H727 cells. This makes imaging of the fibroblasts less complicated than the epithelial cells in tapping mode. Scan sizes differed greatly between Figs. 9 and 10 (89 and 26 µm respectively), as did scan velocity 78 µm/s versus 33 µm/s respectively (0.5 Hz, 0.6 Hz). Gains were set as described in section 2. Unlike the case for contact mode, AC mode image quality was at an optimum when integral gains were set relatively high, 7.2 and 6.3 respectively for figs 9 and 10. In contrast, proportional gains for both images did not differ significantly from contact mode proportional gains, 0.2 and 0.09 for figs 9 and 10 suggesting that integral gain is more influential than proportional gain when imaging live cells. It is unclear as to why the integral gain settings in AC mode needed to be much higher than used in contact mode as scan velocities and size were similar for some of the contact mode images. The natural resonant frequencies in liquid for the 0.32 N/m and 0.58 N/m cantilevers were centred between 10 and 15 kHz respectively. However, driving the cantilevers at these frequencies resulted in no distinguishable images. For this reason the second harmonic for each cantilever was chosen. These were found to be 113 and 100 kHz respectively for the 0.32 N/m and 0.58 N/m cantilever. Shifting to the second harmonic when imaging fibroblasts provided good quality images immediately (fig 9). In contrast, imaging the epithelial cells with the second harmonic provided images of relatively poor quality and only very rarely (fig 10). The behaviour of an oscillating cantilever in liquid is a very complex process. Tipsample interactions are non-linear and anharmonic external forces due to these nonlinear interactions
introduce higher harmonics that excite the system at higher frequencies [Stark and Wolfgang 2003]. It is unclear and beyond the scope of this study to understand why the second harmonics for both cantilevers resulted in images while oscillating at the natural frequencies did not. However it is clear from this study that higher harmonics can be used for imaging live cells and in this case proved to be more useful than using the natural frequencies. It may also be possible to obtain good images of living cells by shifting the frequency to even higher harmonics. References Binnig, G., Quate, C.F (1986) Atomic Force Microscope. Physical Review Letters. 59 (9) 930933 Bushell, G.R., Cahill, C., Clarke, F.M., Gibson, C.T., Myhra, S., Watson, G.S (1999) Analysis of human fibroblasts in vitro-imaging conditions and cytochalasin treatment. Applied Surface Science. 144 141-145 Hansam, H.G., Revenko, I., Kim, K., Laney, D.E (1996) Atomic force microscopy of long and short double-stranded and triple-stranded nucleic acids. Nucleic Acids Research. 24 713-720 Hassan, E.A., Heinz, W.F., Antonik, M.D., D’ Costa, N.P., Nageswaran, S., Schoenenberger, C.A., Hoh, J.H (1998) Relative microelastic mapping of living cells by atomic force microscopy. Biophysical Journal 74 1564-1578 Henderson, E., Haydon, P.G., Sakaguchi, D.S (1992) Actin Filament dynamics in living glial cells imaged by atomic force microscopy. Science. 257 1944-6 Hoh, J.H., Schoenenberger, C.A (1994) Surface morphology and mechanical properties of MDCK monolayers by atomic force microscopy. Journal of Cell Science. 107 1105-1114 Lambrechts, A., Van Troys, M., Ampe, C (2004) The actin cytoskeleton in normal and pathological cell motility. The International Journal of Biochemistry and Cell Biology. 36 1890-1909 Le Grimellec, C., Lesniewska, E., Cachia, C., Schreiber, J.P., Fornel, F.De., Goudonnet, J.P (1994) Imaging of the membrane surface of MDCK cells by atomic force microscopy. Biophysical Journal 67 36-41 Le Grimellec, C., Lesniewska, E., Giocond, M.C., Finot, E., Goudonnet, J.P (1997) Simultaneous imaging of the surface and the submembranous cytoskeleton in living cells by tapping mode atomic force microscopy. C.R Acad.Sci. Paris, Sciences de la vie/life sciences 320 637-643 Le Grimellec, C., Lesniewska, E., Giocond, M.C., Finot, E., Vié, V., Goudonnet, J.P (1998)
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