We thank A. Tager and B. Shea for their ongoing collaboration, and for providing the lung tissue used for demonstration purposes here. This work was supported by National Institutes of Health grant HL-092961. This work was performed in part at the Center for Nanoscale Systems (CNS), a member of the National Nanotechnology Infrastructure Network, which is supported by the National Science Foundation under NSF award ECS-0335765. CNS is part of the Faculty of Arts and Sciences at Harvard University.

1. Lung Tissue Strip Preparation 
<ol>
 <li>Lung tissue is highly compliant and difficult to cut into strips for AFM characterization. To transiently stabilize the lung structure for cutting, inflate isolated mouse lungs intratracheally with 50 ml/kg body weight of 2% low gel point agarose (prepared in PBS) warmed to 37&deg;C. Tie off the trachea and cool the inflated lungs in a bath of PBS at 4&deg;C for 60 minutes. The agarose will gel and stiffen in the airspaces to gently stabilize the lung structure during this interval5. Higher agarose concentrations, i.e. 3-4%, may be used to further enhance tissue stabilization for cutting.</li>
 <li>Cut the agarose-stabilized mouse lung tissue with a razor or scalpel blade into strips of 5 x 5 mm in length and width and 400 &mu;m in thickness, then wash the strips in a PBS bath pre-warmed to 37&deg;C using a 100 ml glass beaker on a bench-top heated stir plate for 5 minutes to remove residual agarose. To exclude large airways and vessels, cut strips from subpleural regions distant from main stem bronchi. If airways and large vessels are to be imaged, cut strips from lung tissue more proximal to main stem bronchi. </li>
</ol>
2. AFM Microindentation and Fluorescence Imaging
<ol>
 <li>To isolate areas of interest for AFM microindentation, tissue can be visualized by phase contrast microscopy, or immunostained and visualized by fluorescence microscopy. For immunostaining, block tissue strips with 5% serum from the source of the desired secondary antibody (in PBS) for 2 hours without fixation or permeabilization at room temperature.</li>
 <li>Incubate the tissue strip with appropriate primary antibody (e.g. rabbit anti-collagen I to visualize interstitial collagen (1:250 dilution), rabbit anti-laminin (1:250 dilution) to visualize basement membrane) in a well of a 12-well plate on a rocker with gentle shaking for 1 hour, wash sample 3 times for 5 minutes each with PBS, followed by appropriate secondary antibody conjugated to fluorescent tag (e.g. Alexa Fluor 546 goat anti-rabbit secondary antibody (1:250 dilution)) for 1 hour at room temperature.</li>
 <li>Wash tissue strips extensively with PBS and store in PBS at 4 &deg;C</li>
 <li>Immediately before AFM characterization, attach the tissue strip to a poly-l-lysine&#x2013;coated 15-mm coverslip by lifting the coverslip from below the floating tissue, making sure the tissue strip spreads evenly on the coverslip surface; if necessary, sandwich the strip with a second clean, uncoated coverslip and apply mild pressure to assist with tissue attachment to the poly-l-lysine&#x2013;coated coverslip. </li>
 <li>Calibrate the AFM system following the manufacture's instruction immediately before each round of microindentation experiments. Determine two critical parameters: 1) the cantilever spring constant using the thermal fluctuation method in air6; and, 2) the cantilever deflection sensitivity, which is a parameter used to scale the photodiode output signal to the actual cantilever deflection distance, &Delta;d. Calibrate deflection sensitivity by obtaining a standard force-displacement curve in PBS on a clean glass slide then calculate the slope of the force-displacement curve (Fig.1B). In the force-displacement curve measurement, the AFM tip is extended towards and retracted from the sample surface at a single location (without rastering in X-Y directions), with the deflection of the cantilever, &Delta;d, monitored as a function of tip displacement, &Delta;z.</li>
</ol>
We recommend using a silicon nitride triangle cantilever with a 5-&mu;m diameter borosilicate spherical tip (Novascan). Using AFM probes with a spring constant of 0.06 N/m, we have mechanically characterized soft materials with shear moduli spanning 100 to 50,000 Pa .
<ol start="6">
 <li>Wipe the bottom surface of the sample coverslip with tissue paper, fasten it to a standard glass slide with vacuum grease, mount the glass slide on the AFM sample stage, and cover the tissue with 500 &mu;l PBS (room temperature). </li>
 <li>Set the microscope for eye-piece viewing to align the AFM tip in the center of the field of view by adjusting the microscope sample stage. Choose an area of interest on the tissue by moving the AFM sample stage, then switch to CCD camera viewing to record phase contrast image and/or fluorescence images of the tissue, as desired. </li>
 <li>Perform standard AFM engagement operation by moving the AFM tip slowly downwards until it is in contact with the sample.</li>
 <li>Accurate AFM microindentation characterization of soft samples requires small indentation depths to avoid large local strains which invalidate the Hertz model used for elastic modulus calculation. To avoid large strains, perform indentation in trigger mode by setting the cantilever maximum deflection (trigger point) to 500 nm. This deflection limit will restrain the maximum indentation force to less than 30 nN (F = cantilever spring constant * displacement, or Fmax = 0.06 nN/nm * 500 nm = 30 nN).</li>
</ol>
The indentation velocity should be selected to be sufficiently slow to explore elastic rather viscoelastic properties of the soft sample7 . A velocity range of 2-20 &mu;m per second is suitable for lung tissue. 
<ol start="10">
 <li>For a single measurement from an area of interest, move the AFM probe to the location of interest and perform a single indentation to collect a standard force-displacement curve (the probe moves only in Z-direction without X-Y scanning).</li>
 <li>For automated mapping of a region of interest, switch to Force-Map mode, select the scan size and sample points within the selected area. In Force-Map mode, the AFM tip rasters across the sample surface between indentation movements, and collects individual force-displacement curves at each point within a defined sample grid. More sampling points will provide higher spatial resolution but lengthen the time required for mapping. We found it practical to use a 16 x 16 sample grid to map an 80 x 80&#x2013;&mu;m area at an indentation velocity of 20 &mu;m per second, which can be completed in ~10 minutes.</li>
</ol>
3. AFM Data Extraction
<ol>
 <li>To calculate the Young's modulus, fit the force-displacement curve to the Hertz spherical indentation model using least squares non-linear curving fitting8 (Fig.1 A,C): 
 <img src="/files/ftp_upload/2911/2911eq1.jpg" alt="Equation 1" /> 
 where F = kc * &Delta;d, is the force to bend the cantilever, kc is the cantilever spring constant, R is sphere tip radius, &delta; = &Delta;z - &Delta;d is indentation and &upsilon; is the sample's 
 Poisson ratio (&upsilon; = 0.4 for lung tissue9). </li>
 <li>To assess the quality of the fit, calculate the SSR value, or the sum of squares of the difference between the data and the fit values, during the non-linear curve fitting. Eliminate unreliable or un-interpretable measurements by discarding data from "bad" curves with large SSR values.</li>
 <li>If desired, convert elastic modulus (E) to shear modulus (G), using the relationship E = 2 x (1 + &upsilon;) x G. To visualize spatial patterns of stiffness collected in Force-Map mode, plot modulus data in contour maps (e.g. in 16 x 16 sample grid covering an 80 x 80&#x2013;&mu;m area).</li>
 <li>To process a large amount of force curve data, a custom algorithm may be written to automatically fit force-displacement curves, extract moduli, and/or plot elastographs using the same procedures and parameters as outlined in 3.2 and 3.3 (e.g. Matlab). </li>
</ol>
<img src="/files/ftp_upload/2911/2911fig1.jpg" alt="Figure 1" /> 
Figure 1 (A) Schematic of AFM microindentation of a soft sample on a rigid substrate using a spherical probe (Reproduced with permission from Dimitriadis E., et al. 2002 Biophys J8). (B) Representative force-displacement curve collected from a clean glass slide in PBS to determine cantilever deflection sensitivity. (C) Representative force-displacement curves collected from soft (blue) and stiff (red) samples showing the corresponding parameters from (A).
4. Representative Results: 
Figure 2A shows a lung parenchyma strip attached on poly-l-lysine coated coverslip in PBS with an AFM probe in place directly above region of interest and ready for microindentation mapping. Figure 2B shows that in properly cut and stained tissue, the alveolar microarchitecture of lung parenchyma is well preserved, as observed by immunofluorescent staining for basement membrane component laminin without fixation or permeabilization. Figure 2 C-D show immunofluorescent staining for collagen I in fresh unfixed lungs harvested from mice previously treated with PBS (Fig. 2C), or treated with bleomycin (Fig. 2D) to induce fibrosis10.
Figure 3A shows sample force-displacement plots obtained from AFM microindentation. With the same applied force, AFM tip generates a large indentation on a soft region (blue line), resulting in a relatively "flat" force-displacement curve versus a small indentation and a "steep" force-displacement curve for a stiffer region (red line). In order to obtain clean force curves, after each indentation the AFM tip needs to be retracted completely from the sample surface and free of contact before the next indentation. This contact free state corresponds to the flat region of the curve in Figure 3A, where the tip translates without cantilever deflection. Figure 3B shows a typical false curve without a flat region which occurs when the tip is not completely retracted from the sample surface (e.g. soft sample attaches to tip) making it impossible to determine the contact point. If the tip is completely trapped in soft sample, the curve may look as shown in Figure 3C, without clear deflection but small noise.
Figure 4 shows shear modulus data extracted from force-displacement curves displayed spatially as stiffness maps (elastographs), where color scales with shear modulus. Elastographs demonstrate striking differences in the range and distribution of tissue shear modulus in normal (Fig. 4A) and fibrotic (Fig. 4B) lung parenchyma, and large spatial variations in shear modulus, particularly within the fibrotic lung sample. 
<img src="/files/ftp_upload/2911/2911fig2.jpg" alt="Figure 2" /> 
Figure 2. (A) Phase contrast micrograph showing an AFM probe over a mouse lung parenchyma strip. Scale bar, 50 &mu;m. (B) Immunostaining of laminin in normal mouse lung tissue showing the alveolar microarchitecture of normal lung parenchyma. White box indicates typical 80-by-80&mu;m elasticity mapping area. Scale bar: 20 &mu;m. (C and D) Immunostaining of collagen I in saline (C) and bleomycin (D) treated mouse lung parenchyma. Scale bars, 100 &mu;m. 
<img src="/files/ftp_upload/2911/2911fig3.jpg" alt="Figure 3" /> 
Figure 3. (A) Representative interpretable force-displacement curves collected from saline (blue) and bleomycin-treated (red) mouse lung parenchyma, respectively. Black lines are the best fit resulting from the spherical Hertz model and are labeled with their corresponding calculated Young's moduli. (B, C) Representative un-interpretable force-displacement curves. 
<img src="/files/ftp_upload/2911/2911fig4.jpg" alt="Figure 4" /> 
Figure 4. Representative elastographs collected from saline (A) and bleomycin-treated (B) mouse lung parenchyma. The color bars indicate shear modulus in kilopascals. Axis labels indicate spatial scale in micrometers. Scale bar: 20 &mu;m

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No conflicts of interest declared.

Mechanical characterization of lung tissue using AFM microindentation offers unprecedented spatial resolution (Fig. 4), providing a unique perspective on microscale variations in tissue stiffness. As an example of its utility, previous macro-scale measurements in normal and fibrotic lung tissue strips indicated an approximate 2-3-fold increase in elastance with fibrosis11,12. In contrast, AFM microindentation reveals that tissue stiffening is highly localized, with some regions exhibiting up to ~30-fold increases in shear modulus above the median observed in normal lung tissue10. As matrix stiffness is now known to critically influence cell function, these local measurements provide invaluable parameters to enhance the biofidelity of cell culture study of lung cells.
Several practical issues arise with the use of thin strips of lung tissue. The surfaces of the strips are not perfectly flat, as the tissue profile follows the architecture of the underlying alveoli. The AFM system automatically adjusts tip position in the Z-direction during indentation when the sample surface height variation is smaller than 15-&mu;m to help overcome this challenge. Measurements are made at room temperature, not 37&deg;C, so deviations in mechanical properties caused by this variation in temperature cannot be evaluated, though they would be expected to be minor. The influence of the underlying alveolar wall architecture on observed mechanical properties is difficult to determine with the current light microscopy setup. For instance, it would be desirable to determine if alveolar walls exhibit anisotropy and different mechanical properties when indented in directions aligned with or transverse to the plane of the wall. However, the current specimens are thick and the imaging system does not have 3D capabilities, hence it is not possible to determine the local alveolar wall orientation at each point of contact. Finally, the influence of cellular constituents on measured mechanical properties remains to be fully elucidated. In the methods detailed here, no efforts are made to specifically remove cellular constituents of the tissue. However, the cells present at the surface available for indentation are unlikely to be viable given the time elapsed since tissue harvest and the necessity of cutting the tissue to gain access to the alveoli. Specific experiments to remove cells, or repopulate matrices with viable cells, and evaluate the resulting changes in tissue stiffness would appear warranted.
Because fresh unfixed tissue is needed for these measurements, the time elapsed from tissue harvest to measurement should be minimized and samples should be stored at 4 &deg;C to avoid changes in mechanical properties. Particular attention should be paid whenever tissue strips are transferred between containers during washing or staining so that minimum distortion or damage is generated. For AFM application in liquid, a crucial step is to cut the tissue as flat as possible and immobilize the sample on the supporting coverslip. If available, an automated sectioning machine such as a vibratome or tissue slicer can be used to cut slices of highly uniform thickness. It's important to attach tissue strips immediately before AFM measurements and minimize the time elapsed for AFM measurements as the sample will eventually detach from the coverslips. One useful observation is that larger strips appear to attach more stably to cover slips and remain in place for longer durations in PBS than smaller strips.
AFM microindentation can characterize samples spanning a broad range from 100 Pa to 50 kPa (shear modulus) when using a standard 0.06 N/m cantilever with a 5 &mu;m diameter spherical tip. This range can be expanded using probes with different spring constants; AFM probes with spherical glass tips ranging from 0.6 to 12 &mu;m in diameter and spring constants ranging from 0.01 to 0.58 N/m are commercially available (e.g. Novascan) and commonly used3. With a 5 &mu;m spherical tip, the theoretical contact area between the tip and tissue is about 5-9 &mu;m2 for 400-700 nm indentation (Fig. 1A). Smaller or larger tips can be used to provide smaller or larger scales of spatial resolution. Pyramidal tips have also been used in AFM microindentation13-16, providing smaller contact areas and thus increasing spatial resolution in mapping, though data fitting is more complex for this tip geometry.
Several limitations to this method should be noted. The lung has traditionally been mechanically characterized non-invasively, for instance using pressure-volume analysis17 or punch-indentation of whole lungs19,20. Invasive methods such as the one described here alter the lung architecture in important ways through the loss of the air-liquid interface that normally exists in the air-filled lung and the loss of pre-stress that maintains lung partial inflation upon relaxation of respiratory muscles. These limitations are common to all measurements made in lung tissue strips18. Notably, however, the median stiffness measured in the parenchyma of normal lung tissue (shear modulus ~0.5kPa) does not differ substantially from estimates based on punch-indentation of intact lungs at resting volumes19,20. While lung tissue is known to exhibit non-linear stiffening with increasing deformation, it is not possible to test in a rigorous fashion whether this property persists down to the micro-scale with the methods employed here. The Hertz model assumes homogeneity of the sample. However, most biological materials, including lung parenchyma, are increasingly heterogeneous at decreasing spatial scales. Heterogeneity of the sample can result in artifacts like variation of the Young's modulus depending on indentation depth, i.e. depending on the layer or component that is deforming. The heterogeneity in the xy-plane can be limited by carefully choosing the appropriate spherical tip size depending on the microstructure of the biomaterial as proposed by Dimitriadis EK et al.8 It is much more difficult to predict or correct the Hertz model error due to material heterogeneity in the z-direction. Azeloglu et al. recently proposed a hybrid computational model to characterize the elastic properties of heterogeneous substrate with discrete embedded inclusions21. Their new technique provides a potential means to calculate inclusion properties of heterogeneous materials overcoming the limitations of Hertzian analysis. 
The Hertz model also assumes absolute elastic behavior, while biological materials typically display time-dependent viscoelastic behaviors. A full viscoelastic characterization of tissue can be obtained by varying the indentation velocities used. Importantly, previous macro-scale mechanical testing of normal and fibrotic lung tissue demonstrates weak frequency dependence of lung tissue mechanical properties, and preservation of the differences between normal and fibrotic tissue mechanical properties across all frequencies tested11. These findings strongly suggest that the measurement of mechanical properties using a single indentation velocity with AFM captures an essential aspect of the changes in tissue mechanical properties that accompany fibrosis. 
The Poisson's ratio of 0.4 for lung tissue used in this work is from a macroscopic measurement9. Unfortunately, the Poisson's ratio at micro-scale and any changes under disease condition are not available in the literature. As alternatives to E, E/(1-&upsilon;2) or (1-&upsilon;2)/&pi;E (denoted the elastic constant k)22 can be calculated from AFM microindentation and reported when the Poisson's ratio is unknown. For most biomaterials Poisson's ratio is in the range of 0.4 to 0.5 due to their high water content. Within the range 0.3-0.5, the factor 1/(1-&upsilon;2) varies only from 1.10-1.33, such that reasonable variations in the Poisson's ratio exert only modest effects on the reported modulus. The increase in shear modulus that we report for fibrotic tissue relative to normal tissue is several fold in magnitude, implying that errors associated with variations in Poisson's ratio are minor relative to the changes in mechanical properties observed. 
The actual algorithm and code that can be used for the analysis of force-displacement data is subject to the specific application condition and the subsequent characteristics of the diverse populations of force-displacement curves. If more sophisticated analysis is of interest, one may consult the work of Lin et al.23. The authors compiled a series of synergistic strategies into an algorithm that overcomes many of the complications that have previously impeded efforts to automate the fitting of Hertz models of indentation data. 
Several other areas are available for further development and exploitation of this method. In cases where one is interested in visualizing the alveolar walls without antibody labeling, both elastin and collagen can be visualized from their autofluorescent signal in the green spectrum. On the other hand, better imaging, using either thinner tissue sections, 3D imaging techniques, or both, could enhance the ability to correlate tissue architecture with underlying mechanical properties. While the current methods allow staining and visualization of extracellular matrix components such as collagen and laminin, additional efforts could be aimed at staining cell surface markers to identify specific cell populations and to characterize the mechanical microenvironments in the vicinity of such populations. Alternatively, tissue could be harvested from mice expressing fluorescently-tagged lineage markers or cell specific proteins to pursue the same goal. Finally, the method detailed here appears well suited to characterizing other anatomical features in the lung, such as vessels which remodel in hypertension, and airways which remodel in asthma. Based on its current state of development and potential for further enhancement, AFM microindentation appears poised to yield valuable insights into the changes in tissue stiffness that accompany disease progression in the lung, and will no doubt be of value in characterizing spatial and temporal changes in the stiffness of a variety of other soft tissues.

<table><tbody><tr><td>AFM tip</td><td>Novascan Technologies, Inc.</td><td>PT.GS</td><td>5 micron Borosilicate bead, 0.06 N/m</td></tr><tr><td>Poly-L-Lysine coverslips</td><td>VWR international</td><td>354085</td><td>BD BioCoat 12 mm round No.1 glass</td></tr><tr><td>Agarose, Low Gelling Temperature</td><td>Sigma-Aldrich</td><td>A0701</td><td></td></tr><tr><td>Rabbit anti-Collagen I (Rb pAb) antibody</td><td>Abcam</td><td>ab34710-100</td><td></td></tr><tr><td>Alexa Fluor 546 goat anti-rabbit antibody </td><td>Invitrogen</td><td>A11010</td><td></td></tr><tr><td>Rabbit anti-Laminin </td><td>Sigma-Aldrich</td><td>L9393</td><td></td></tr></tbody></table>

micro-mechanical characterization of lung tissue using atomic force microscopy

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3. On the macro scale the stiffness of tissues and organs within the human body span several orders of magnitude4. Much less is known about how stiffness varies spatially within tissues, and what the scope and spatial scale of stiffness changes are in disease processes that result in tissue remodeling. To better understand how changes in matrix stiffness contribute to cellular physiology in health and disease, measurements of tissue stiffness obtained at a spatial scale relevant to resident cells are needed. This is particularly true for the lung, a highly compliant and elastic tissue in which matrix remodeling is a prominent feature in diseases such as asthma, emphysema, hypertension and fibrosis. To characterize the local mechanical environment of lung parenchyma at a spatial scale relevant to resident cells, we have developed methods to directly measure the local elastic properties of fresh murine lung tissue using atomic force microscopy (AFM) microindentation. With appropriate choice of AFM indentor, cantilever, and indentation depth, these methods allow measurements of local tissue shear modulus in parallel with phase contrast and fluorescence imaging of the region of interest. Systematic sampling of tissue strips provides maps of tissue mechanical properties that reveal local spatial variations in shear modulus. Correlations between mechanical properties and underlying anatomical and pathological features illustrate how stiffness varies with matrix deposition in fibrosis. These methods can be extended to other soft tissues and disease processes to reveal how local tissue mechanical properties vary across space and disease progression.

Watch this Scientific Journal Video about Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy at JoVE.com

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

The stiffness of the extracellular matrix strongly influences multiple behaviors of adherent cells. Matrix stiffness varies spatially throughout a tissue, and undergoes modification in various disease conditions. Here we develop methods to characterize spatial variations in stiffness in normal and fibrotic mouse lung tissue using atomic force microscopy microindentation.

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3.  On the macro scale the stiffness of tissues and organs ...

micro-mechanical-characterization-lung-tissue-using-atomic-force

Nanyang Technological University

Research

JoVE Journal

Biology

22.8K Views.  Harvard School of Public Health. The stiffness of the extracellular matrix strongly influences multiple behaviors of adherent cells. Matrix stiffness varies spatially throughout a tissue, and undergoes modification in various disease conditions. Here we develop methods to characterize spatial variations in stiffness in normal and fibrotic mouse lung tissue using atomic force microscopy microindentation.

Video: Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be measured to ~10 pN by a laser and sectored detector, which can be converted to image topography. Amplitude modulation or &#8220;tapping mode&#8221; AFM involves the probe making intermittent contact with the surface while oscillating at its resonant frequency to produce an image. Used in conjunction with a fluid cell, tapping-mode AFM enables the imaging of biological macromolecules such as proteins in physiologically relevant conditions. Tapping-mode AFM requires manual tuning of the probe and frequent adjustments of a multitude of scanning parameters which can be challenging for inexperienced users. To obtain high-quality images, these adjustments are the most time consuming.
PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) produces an image by measuring a force response curve for every point of contact with the sample. With ScanAsyst software, PF-QNM can be automated. This software adjusts the set-point, drive frequency, scan rate, gains, and other important scanning parameters automatically for a given sample. Not only does this process protect both fragile probes and samples, it significantly reduces the time required to obtain high resolution images. PF-QNM is compatible for AFM imaging in fluid; therefore, it has extensive application for imaging biologically relevant materials.
The method presented in this paper describes the application of PF-QNM to obtain images of a bacterial red-light photoreceptor, RpBphP3 (P3), from photosynthetic R. palustris in its light-adapted state. Using this method, individual protein dimers of P3 and aggregates of dimers have been observed on a mica surface in the presence of an imaging buffer. With appropriate adjustments to surface and/or solution concentration, this method may be generally applied to other biologically relevant macromolecules and soft materials.

A method for investigating the structure of a protein photoreceptor using atomic force microscopy (AFM) is described in this paper. PeakForce Quantitative Nanomechanical Property Mapping (PF-QNM) reveals intact protein dimers on a mica surface.

Atomic force microscopy (AFM) uses a pyramidal tip attached to a cantilever to probe the force response of a surface. The deflections of the tip can be ...

Engineering

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Bioengineering

Sub-nanometer Resolution Imaging with Amplitude-modulation Atomic Force Microscopy in Liquid

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown that when operated in amplitude-modulation (tapping) mode, atomic or molecular-level resolution images can be achieved over a wide range of soft and hard samples in liquid. In these situations, small oscillation amplitudes (SAM-AFM) enhance the resolution by exploiting the solvated liquid at the surface of the sample. Although the technique has been successfully applied across fields as diverse as materials science, biology and biophysics and surface chemistry, obtaining high-resolution images in liquid can still remain challenging for novice users. This is partly due to the large number of variables to control and optimize such as the choice of cantilever, the sample preparation, and the correct manipulation of the imaging parameters. Here, we present a protocol for achieving high-resolution images of hard and soft samples in fluid using SAM-AFM on a commercial instrument. Our goal is to provide a step-by-step practical guide to achieving high-resolution images, including the cleaning and preparation of the apparatus and the sample, the choice of cantilever and optimization of the imaging parameters. For each step, we explain the scientific rationale behind our choices to facilitate the adaptation of the methodology to every user's specific system.

We present a method for achieving sub-nanometer resolution images with amplitude-modulation (tapping mode) atomic force microscopy in liquid. The method is demonstrated on commercial atomic force microscopes. We explain the rationale behind our choices of parameters and suggest strategies for resolution optimization.

Atomic force microscopy (AFM) has become a well-established technique for nanoscale imaging of samples in air and in liquid. Recent studies have shown ...

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in indentation mode, we show how to measure an elastic modulus and use it to evaluate cell wall mechanical properties. In addition, we also explain how to evaluate turgor pressure. The main advantages of atomic force microscopy are that it is non-invasive, relatively rapid (5~20 min), and that virtually any type of living plant tissue that is superficially flat can be analyzed without the need for treatment. The resolution can be very good, depending on the tip size and on the number of measurements per unit area. One limitation of this method is that it only gives direct access to the superficial cell layer.

Here, we present atomic force microscopy (AFM), operated as a nano- and micro-indentation tool on cells and tissues. The instrument allows the simultaneous acquisition of 3D surface topography of the sample and its mechanical properties, including cell wall Young's modulus as well as turgor pressure.

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in ...

Developmental Biology

Atomic Force Microscopy Cantilever-Based Nanoindentation: Mechanical Property Measurements at the Nanoscale in Air and Fluid

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by the probe tip and its contact area with the sample can be quantified, it is possible to determine the nanoscale mechanical properties (e.g., elastic or Young&#39;s modulus) of the surface being probed. A detailed procedure for performing quantitative AFM cantilever-based nanoindentation experiments is provided here, with representative examples of how the technique can be applied to determine the elastic moduli of a wide variety of sample types, ranging from kPa to GPa. These include live mesenchymal stem cells (MSCs) and nuclei in physiological buffer, resin-embedded dehydrated loblolly pine cross-sections, and Bakken shales of varying composition.
Additionally, AFM cantilever-based nanoindentation is used to probe the rupture strength (i.e., breakthrough force) of phospholipid bilayers. Important practical considerations such as method choice and development, probe selection and calibration, region of interest identification, sample heterogeneity, feature size and aspect ratio, tip wear, surface roughness, and data analysis and measurement statistics are discussed to aid proper implementation of the technique. Finally, co-localization of AFM-derived nanomechanical maps with electron microscopy techniques that provide additional information regarding elemental composition is demonstrated.

Quantifying the contact area and force applied by an atomic force microscope (AFM) probe tip to a sample surface enables nanoscale mechanical property determination. Best practices to implement AFM cantilever-based nanoindentation in air or fluid on soft and hard samples to measure elastic modulus or other nanomechanical properties are discussed.

An atomic force microscope (AFM) fundamentally measures the interaction between a nanoscale AFM probe tip and the sample surface. If the force applied by ...

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record forces curves on 1000 cells in 4 hours automatically. To maintain a 4 hour analysis time, the number of force curves per cell is reduced to 9 or 16. The method combines a Jython based program and a strategy for assembling cells on defined patterns. The program, implemented on a commercial Bio-AFM, can center the tip on the first cell of the array and then move, automatically, from cell to cell while recording force curves on each cell. Using this methodology, it is possible to access the biophysical parameters of the cells such as their rigidity, their adhesive properties, etc. With the automation and the large number of cells analyzed, one can access the behavior of the cell population. This is a breakthrough in the Bio-AFM field where data have, so far, been recorded on only a few tens of cells.

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

This protocol aims to automate AFM measurements on hundreds of microbial cells. First, microbes are immobilized into PDMS stamp microstructures and then force spectroscopy measurements are performed automatically on hundreds of immobilized cells.

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record ...

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning and transmission electron microscope, to the invention of atomic force microscopy, and advances in fluorescent imaging, there have been substantial gains in technologies that enable the study of small materials. Conpokal is a portmanteau that combines confocal microscopy with atomic force microscopy (AFM), where a probe &#8220;pokes&#8221; the surface. Although each technique is extremely effective for the qualitative and/or quantitative image collection on their own, Conpokal provides the capability to test with blended fluorescence imaging and mechanical characterization. Designed for near simultaneous confocal imaging and atomic force probing, Conpokal facilitates experimentation on live microbiological samples. The added insight from paired instrumentation provides co-localization of measured mechanical properties (e.g., elastic modulus, adhesion, surface roughness) by AFM with subcellular components or activity observable through confocal microscopy. This work provides a step by step protocol for the operation of laser scanning confocal and atomic force microscopy, simultaneously, to achieve same cell, same region, confocal imaging, and mechanical characterization.

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy Conpokal on Live Cells

Presented here is the protocol of the Conpokal technique, which combines confocal and atomic force microscopy into a single instrument platform. Conpokal provides same cell, same region, near simultaneous confocal imaging and mechanical characterization of live biological samples.

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning ...

Addressing Practical Issues in Atomic Force Microscopy-Based Micro-Indentation on Human Articular Cartilage Explants

Without a doubt, atomic force microscopy (AFM) is currently one of the most powerful and useful techniques to assess micro and even nano-cues in the biological field. However, as with any other microscopic approach, methodological challenges can arise. In particular, the characteristics of the sample, sample preparation, type of instrument, and indentation probe can lead to unwanted artifacts. In this protocol, we exemplify these emerging issues on healthy as well as osteoarthritic articular cartilage explants. To this end, we first show via a step-by-step approach how to generate, grade, and visually classify ex vivo articular cartilage discs according to different stages of degeneration by means of large 2D mosaic fluorescence imaging of the whole tissue explants. The major strength of the ex vivo model is that it comprises aged, native, human cartilage that allows the investigation of osteoarthritis-related changes from early onset to progression. In addition, common pitfalls in tissue preparation, as well as the actual AFM procedure together with the subsequent data analysis, are also presented. We show how basic but crucial steps such as sample preparation and processing, topographic sample characteristics caused by advanced degeneration, and sample-tip interaction can impact data acquisition. We also subject to scrutiny the most common problems in AFM and describe, where possible, how to overcome them. Knowledge of these limitations is of the utmost importance for correct data acquisition, interpretation, and, ultimately, the embedding of findings into a broad scientific context.

We present a step-by-step approach to identify and address the most common problems associated with atomic force microscopy micro-indentations. We exemplify the emerging problems on native human articular cartilage explants characterized by various degrees of osteoarthritis-driven degeneration.

Without a doubt, atomic force microscopy (AFM) is currently one of the most powerful and useful techniques to assess micro and even nano-cues in the ...

Extracting the Young's Modulus of Native Murine Pulmonary Basement Membranes from Atomic Force Microscopy Derived Force Maps

Atomic force microscopy (AFM) allows the characterization of the mechanical properties of a sample with a spatial resolution of several tens of nanometers. Because mammalian cells sense and react to the mechanics of their immediate microenvironment, the characterization of biomechanical properties of tissues with high spatial resolution is crucial for understanding various developmental, homeostatic, and pathological processes. The basement membrane (BM), a roughly 100 - 400 nm thin extracellular matrix (ECM) substructure, plays a significant role in tumor progression and metastasis formation. Although determining Young&#39;s modulus of such a thin ECM substructure is challenging, biomechanical data of the BM provides fundamental new insights into how the BM affects cell behavior and, in addition, offers valuable diagnostic potential. Here, we present a visualized protocol for assessing BM mechanics in murine lung tissue, which is one of the major organs prone to metastasis. We describe an efficient workflow for determining the Young&#39;s modulus of the BM, which is located between the endothelial and epithelial cell layers in lung tissue. The step-by-step instructions comprise murine lung tissue freezing, cryosectioning, and AFM force-map recording on tissue sections. Additionally, we provide a semi-automatic data analysis procedure using the CANTER Processing Toolbox, an in-house developed user-friendly AFM data analysis software. This tool enables automatic loading of recorded force maps, conversion of force versus piezo-extension curves to force versus indentation curves, computation of Young&#39;s moduli, and generation of Young&#39;s modulus maps. Finally, it shows how to determine and isolate Young&#39;s modulus values derived from the pulmonary BM through the use of a spatial filtering tool.

This protocol visualizes how to prepare cryosections of murine lung tissue, perform atomic force microscopy force map experiments, and analyze the data to determine Young's modulus of the native murine pulmonary basement membrane.

Atomic force microscopy (AFM) allows the characterization of the mechanical properties of a sample with a spatial resolution of several tens of ...

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell behavior such as cell function, differentiation, and motility. However, as these processes are not homogeneous throughout the whole organ, it has become increasingly important to understand changes in the mechanical properties of tissues on the cellular level.
To be able to monitor the stiffening of collagen-rich areas within the liver lobes, this paper presents a protocol for measuring liver tissue elastic moduli with high spatial precision by atomic force microscopy (AFM). AFM is a sensitive method with the potential to characterize local mechanical properties, calculated as Young&#39;s (also referred to as elastic) modulus. AFM coupled with polarization microscopy can be used to specifically locate the areas of fibrosis development based on the birefringence of collagen fibers in tissues. Using the presented protocol, we characterized the stiffness of collagen-rich areas from fibrotic mouse livers and corresponding areas in the livers of control mice.
A prominent increase in the stiffness of collagen-positive areas was observed with fibrosis development. The presented protocol allows for a highly reproducible method of AFM measurement, due to the use of mildly fixed liver tissue, that can be used to further the understanding of disease-initiated changes in local tissue mechanical properties and their effect on the fate of neighboring cells.

We present a protocol to measure the elastic moduli of collagen-rich areas in normal and diseased liver using atomic force microscopy. The simultaneous use of polarization microscopy provides high spatial precision for localizing collagen-rich areas in the liver sections.

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell ...

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Summary

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Chapters in this video

Atomic Force Microscopy of Red-Light Photoreceptors Using PeakForce Quantitative Nanomechanical Property Mapping

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Sub-nanometer Resolution Imaging with Amplitude-modulation Atomic Force Microscopy in Liquid

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

Atomic Force Microscopy Cantilever-Based Nanoindentation: Mechanical Property Measurements at the Nanoscale in Air and Fluid

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

Addressing Practical Issues in Atomic Force Microscopy-Based Micro-Indentation on Human Articular Cartilage Explants

Extracting the Young's Modulus of Native Murine Pulmonary Basement Membranes from Atomic Force Microscopy Derived Force Maps

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy