Name: micro-mechanical characterization of lung tissue using atomic force microscopy
Uploaded: 2011-08-28T14:00:00
Duration: 11 min 10 s
Description: Watch this Scientific Journal Video about Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy at JoVE.com

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.

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			<th>Catalogue number</th>
			<th>Comments</th>
		</tr>
		<tr>
			<td>AFM tip</td>
			<td>Novascan</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</td>
			<td>354085</td>
			<td>BD BioCoat 12 mm round No.1 glass</td>
		</tr>
		<tr>
			<td>Agarose, Low Gelling Temperature</td>
			<td>Sigma</td>
			<td>A0701</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rabbit anti-Collagen I (Rb pAb) antibody</td>
			<td>Abcam</td>
			<td>ab34710-100</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Alexa Fluor 546 goat anti-rabbit antibody </td>
			<td>Invitrogen</td>
			<td>A11010</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rabbit anti-Laminin </td>
			<td>Sigma</td>
			<td>L9393</td>
			<td>&nbsp;</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

Research

JoVE Journal

Biology

Live Cell Response to Mechanical Stimulation Studied by Integrated Optical and Atomic Force Microscopy

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to investigate cells behavior at sub-cellular level with high spatial and temporal resolution was developed. Thus, an atomic force microscope (AFM) was integrated with total internal reflection fluorescence (TIRF) microscopy and fast-spinning disk (FSD) confocal microscopy. The integrated system is broadly applicable across a wide range of molecular dynamic studies in any adherent live cells, allowing direct optical imaging of cell responses to mechanical stimulation in real-time. Significant rearrangement of the actin filaments and focal adhesions was shown due to local mechanical stimulation at the apical cell surface that induced changes into the cellular structure throughout the cell body. These innovative techniques will provide new information for understanding live cell restructuring and dynamics in response to mechanical force. A detailed protocol and a representative data set that show live cell response to mechanical stimulation are presented.

This paper aims to instruct the reader in the operation of an integrated atomic force-optical imaging microscope for mechanical stimulation of live cells in culture. A step-by-step protocol is presented. A representative data set that shows live cell response to mechanical stimulation is presented.

To understand the mechanism by which living cells sense mechanical forces, and how they respond and adapt to their environment, a new technology able to ...

Identifying Targets of Human microRNAs with the LightSwitch Luciferase Assay System using 3'UTR-reporter Constructs and a microRNA Mimic in Adherent Cells

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes. More than 700 human miRNAs have been identified so far with each having up to hundreds of unique target mRNAs. Computational tools, expression and proteomics assays, and chromatin-immunoprecipitation-based techniques provide important clues for identifying mRNAs that are direct targets of a particular miRNA. In addition, 3'UTR-reporter assays have become an important component of thorough miRNA target studies because they provide functional evidence for and quantitate the effects of specific miRNA-3'UTR interactions in a cell-based system. To enable more researchers to leverage 3'UTR-reporter assays and to support the scale-up of such assays to high-throughput levels, we have created a genome-wide collection of human 3'UTR luciferase reporters in the highly-optimized LightSwitch Luciferase Assay System. The system also includes synthetic miRNA target reporter constructs for use as positive controls, various endogenous 3'UTR reporter constructs, and a series of standardized experimental protocols.
Here we describe a method for co-transfection of individual 3'UTR-reporter constructs along with a miRNA mimic that is efficient, reproducible, and amenable to high-throughput analysis.

Identifying Targets of Human microRNAs with the LightSwitch Luciferase Assay System using 3&#039;UTR-reporter Constructs and a microRNA Mimic in Adherent Cells

MicroRNAs (miRNAs) are important regulators of gene expression and have been shown to play a role in numerous biological processes. To better understand miRNA-UTR interactions, we have created a genome-wide collection of 3 UTR luciferase reporters paired with a novel luciferase gene and assay reagent, the LightSwitch system.

MicroRNAs (miRNAs) are important regulators of gene expression and play a role in many biological processes.  More than 700 human miRNAs have been ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Characterizing Mechanical Properties of Primary Cell Wall in Living Plant Organs Using Atomic Force Microscopy

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the plant. Many sophisticated techniques have been developed to measure these properties; however, atomic force microscopy (AFM) remains the most convenient for studying cell wall elasticity at the cellular level. One of the most important limitations of this technique has been that only superficial or isolated living cells can be studied. Here, the use of atomic force microscopy to investigate the mechanical properties of primary cell walls belonging to the internal tissues of a plant body is presented. This protocol describes measurements of the apparent Young&#39;s modulus of cell walls in roots, but the method can also be applied to other plant organs. The measurements are performed on vibratome-derived sections of plant material in a liquid cell, which allows (i) avoiding the use of plasmolyzing solutions or sample impregnation with wax or resin, (ii) making the experiments fast, and (iii) preventing dehydration of the sample. Both anticlinal and periclinal cell walls can be studied, depending on how the specimen was sectioned. Differences in the mechanical properties of different tissues can be investigated in a single section. The protocol describes the principles of study planning, issues with specimen preparation and measurements, as well as the method of selecting force-deformation curves to avoid the influence of topography on the obtained values of elastic modulus. The method is not limited by sample size but is sensitive to cell size (i.e., cells with a large lumen are difficult to examine).

Studies of cell wall biomechanics are essential for understanding plant growth and morphogenesis. The following protocol is proposed to investigate thin primary cell walls in the internal tissues of young plant organs using atomic force microscopy.

The mechanical properties of the primary cell walls determine the direction and rate of plant cell growth and, therefore, the future size and shape of the ...

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 ...

Contact Mode Atomic Force Microscopy as a Rapid Technique for Morphological Observation and Bacterial Cell Damage Analysis

Electron microscopy is one of the tools required to characterize cellular structures. However, the procedure is complicated and expensive due to the sample preparation for observation. Atomic force microscopy (AFM) is a very useful characterization technique due to its high resolution in three dimensions and because of the absence of any requirement for vacuum and sample conductivity. AFM can image a wide variety of samples with different topographies and different types of materials. 
AFM provides high-resolution 3D topography information from the angstrom level to the micron scale. Unlike traditional microscopy, AFM uses a probe to generate an image of the surface topography of a sample. In this protocol, the use of this type of microscopy is suggested for the morphological and cell damage characterization of bacteria fixed on a support. Strains of Staphylococcus aureus (ATCC 25923), Escherichia coli (ATCC 25922), and Pseudomonas hunanensis (isolated from garlic bulb samples) were used. In this work, bacterial cells were grown in specific culture media. To observe cell damage, Staphylococcus aureus and Escherichia coli were incubated with different concentrations of nanoparticles (NPs). 
A drop of bacterial suspension was fixed on a glass support, and images were taken with AFM at different scales. The images obtained showed the morphological characteristics of the bacteria. Further, employing AFM, it was possible to observe the damage to the cellular structure caused by the effect of NPs. Based on the images obtained, contact AFM can be used to characterize the morphology of bacterial cells fixed on a support. AFM is also a suitable tool for the investigation of the effects of NPs on bacteria. Compared to electron microscopy, AFM is an inexpensive and easy-to-use technique.

Here, we present the application of atomic force microscopy (AFM) as a simple and fast method for bacterial characterization and analyze details such as the bacterial size and shape, bacterial culture biofilms, and the activity of nanoparticles as bactericides.

Electron microscopy is one of the tools required to characterize cellular structures. However, the procedure is complicated and expensive due to the ...

Measurement of Stiffness in Mouse Retinal Capillaries and Subendothelial Matrix

Isolation of Mouse Retinal Capillaries and Subendothelial Matrix for Stiffness Measurement Using Atomic Force Microscopy

Retinal capillary degeneration is a clinical hallmark of the early stages of diabetic retinopathy (DR). Our recent studies have revealed that diabetes-induced retinal capillary stiffening plays a crucial and previously unrecognized causal role in inflammation-mediated degeneration of retinal capillaries. The increase in retinal capillary stiffness results from the overexpression of lysyl oxidase, an enzyme that crosslinks and stiffens the subendothelial matrix. Since tackling DR at the early stage is expected to prevent or slow down DR progression and associated vision loss, subendothelial matrix, and capillary stiffness represent relevant and novel therapeutic targets for early DR management. Further, direct measurement of retinal capillary stiffness can serve as a crucial preclinical validation step for the development of new imaging techniques for non-invasive assessment of retinal capillary stiffness in animal and human subjects. With this view in mind, we here provide a detailed protocol for the isolation and stiffness measurement of mouse retinal capillaries and subendothelial matrix using atomic force microscopy.

Author Spotlight: Investigating Vascular Stiffness and Inflammation in Early Diabetic Retinopathy

We recently identified retinal capillary stiffening as a new paradigm for retinal dysfunction associated with diabetes. This protocol elaborates the steps for isolation of mouse retinal capillaries and the subendothelial matrix from retinal endothelial cultures, followed by a description of the stiffness measurement technique using atomic force microscopy.

Retinal capillary degeneration is a clinical hallmark of the early stages of diabetic retinopathy (DR). Our recent studies have revealed that ...

Mechanical Characterization of Tissue Stiffness During Axolotl Limb Regeneration

Atomic Force Microscopy Measurements of Cartilage in Intact and Regenerating Axolotl Limbs

Mechanical forces provide important signals for normal cell function and pattern formation in developing tissues, and their role has been widely studied during embryogenesis and pathogenesis. Comparatively, little is known of these signals during animal regeneration. 
The axolotl is an important model organism for the study of regeneration, given its ability to fully restore many organs and tissues after injury, including missing cartilage and bone. Due to its crucial role as the main supporting tissue in the vertebrate body, regaining skeletal function during regeneration requires both the restoration of the missing structures as well as their mechanical properties. This protocol describes a method for processing axolotl limb samples for atomic force microscopy (AFM), which is the gold standard for probing cell and tissue mechanical properties at high spatial resolution. 
Taking advantage of the regenerative capabilities of the axolotl, this study measured the stiffness of limb cartilage during homeostasis and two stages of limb regeneration: tissue histolysis and cartilage condensation. We show that AFM is a valuable tool for gaining insights into dynamic tissue restructuring and the mechanical changes that occur during regeneration.

Author Spotlight: Exploring the Role of Mechanical Signals in Tissue Regeneration Through Atomic Force Microscopy 

In this protocol, we show how to prepare axolotl tissue for atomic force microscopy (AFM) and perform indentation measurements in intact and regenerating limb cartilage.

Mechanical forces provide important signals for normal cell function and pattern formation in developing tissues, and their role has been widely studied ...

Axolotl Tissue Mounting and Processing for Atomic Force Microscopy Measurements in Regenerating and Intact Limb Tissues

To prepare one-centimeter long cylinders of one centimeter diameter, heat a cardboard cutter under a Bunsen burner flame, and cut a 15-milliliter tube with the heated blade. Cover one side of the cylinders with Parafilm pieces. After anesthethetizing the axolotl and making sure it is unresponsive to tactile stimuli, orient the limb perpendicular to the body axis.Amputate the limb immediately distal to the calcified area of the zeugopodial region. Return the axolotl to a tank containing fresh holding water with analgesics. After 24 hours, transfer the axolotl back to fresh holding water and allow the limb to regenerate until the desired stage of interest.After placing the anesthetized axolotl under a stereo microscope, measure the length of the limb at the desired stage of regeneration for histolysis and condensing cartilage stages. After collecting the limbs, place them in a Petri dish for dissection. Under a stereo microscope, dissect the limbs with a scalpel at the elbow level, and place them in a Petri dish containing APBS solution.Arrange the Pasteur pipettes and the thermoblock set at 37 degrees Celsius with agarose aliquots on the working platform. Remove the cold block from the minus 20 degrees Celsius freezer and place the cylinder with the Parafilm-covered end facing down on top of it. Place the limb on a clean plate and gently remove excess liquid from the dissected limb with tissue paper.Add the melted low melting point agarose on top, and briefly move the limb around in the agarose to displace any remaining APBS from the skin surface. Quickly place the limb inside the cylinder, ensuring it is oriented vertically with the area of interest facing upwards. Add low melting point agarose inside the cylinder until the tissue is fully covered.Next, remove the cylinder from the cold block and allow the agarose to solidify for 30 seconds. After solidification, remove the Parafilm from the bottom of the cylinder and attach the agarose-containing tissue to the vibratome stage with cyanoacrylate glue. Then, remove any excess surrounding agarose with a scalpel.Submerge the stage in APBS solution for sectioning. Begin sectioning the agarose in 100-micrometer steps until the tissue tip is reached. Then, section until the distal portion of tissue is removed.Using a razor blade, carefully remove the tissue-containing agarose block from the vibratome stage. Immediately, attach the block to a 35-millimeter plastic Petri dish with surgical tissue adhesive glue. Add approximately two milliliters of culture medium to cover the tissue.