This work was supported by the Grant Agency of the Czech Republic (18-02699S), the Institutional Research Project of the Czech Academy of Sciences (RVO 68378050), and MEYS CR project NICR EXCELES (LX22NPO05102). CIISB, Instruct-CZ Centre of Instruct-ERIC EU consortium, funded by MEYS CR infrastructure project LM2018127 and European Regional Development Fund-Project &#34;UP CIISB&#34; (No. CZ.02.1.01/0.0/0.0/18_046/0015974) financially supported the measurements at the CF Nanobiotechnology, CEITEC MU. We also acknowledge the Light Microscopy Core Facility, IMG CAS, Prague, Czech Republic, supported by MEYS (LM2018129, CZ.02.1.01/0.0/0.0/18_046/0016045) and RVO: 68378050-KAV-NPUI, for their support with the microscopy imaging presented herein.

The authors have no conflicts of interest to disclose.

The presented protocol provides a step-by-step reproducible method for AFM measurement of normal and fibrotic mouse liver tissue. Coupled polarization microscopy provides high spatial precision and enables visualization of collagen fibers due to their birefringence. Further, a detailed description of the analysis of the obtained force curves is provided. AFM stiffness measurement can be performed on the cellular level, which allows local changes in liver tissue mechanical properties due to developing fibrotic disease to be monitored. Liver fibrosis is not a homogeneous process affecting the entire organ. On the contrary, areas of collagen-rich fibrotic septa are interspersed with areas of low or no collagen deposits. Thus, stiffness changes are specific to the local microenvironment and only affect cells locally in contact with areas damaged by injury. This microscale of stiffness heterogeneity is also apparent in the details of AFM Young&#39;s modulus maps, where points of high stiffness border the areas of almost normal stiffness. This variation shows that even collagen scar tissue area is not mechanically homogeneous and requires AFM measurement to be characterized on a cellular level (Figure 4).
The presented protocol allows the measurements of liver stiffness by AFM independently of the liver collection, as the whole liver lobes embedded in OCT can be stored for a prolonged period at &#8722;80 &#176;C. However, once the tissue is sectioned, we recommend measuring the samples within ~2 weeks as we have observed a gradual softening of tissue sections stored for longer periods of time (Figure 5).
The AFM equipped with polarization microscopy allows for precisely locating the area of interest within the liver lobule structure. However, it also has some limitations that need to be considered when interpreting the results. The stiffness values obtained here were measured at room temperature. We assume that the effects of temperature on the mechanical properties of soft tissue will be small; however, this might be one of the reasons for differences between the reported in vivo values of mechanical properties of liver tissues and the values in this study.
Furthermore, this protocol allows AFM analysis of liver tissue for up to 3 h, which requires mild fixation of the tissue. The mild fixation of tissue sections, as well as the freeze-thaw cycle, will most likely affect the absolute values of Young&#39;s modulus. Thus, the reported values of Young&#39;s moduli might differ from in vivo values. Further studies are needed to optimize the protocol for the measurement of absolute values of Young&#39;s modulus from liver sections, which may be achieved by a different method for fixation of liver tissue64.
Nonetheless, we observed increasing stiffness of collagen-rich areas in the livers of mice treated with CCl4 for 3 weeks compared to 6 weeks. Such changes correspond to fibrosis progression during prolonged injury (Figure 4) and show that relative differences can be probed between different treatments using the presented protocol. This is in agreement with the observations of Cal&#242; et al.,&#160;who showed that mildly fixed liver sections show similar differences in stiffness values between collagen-rich and collagen-lacking areas as in fresh tissue25.
We used the SD-qp-BioT-TL-10&#160;cantilever (theoretical spring constant ~0.09 N/m) modified with a 5.7 &#181;m diameter spherical tip to minimize mechanical disruption of the liver tissue during measurements. A 5.7 &#181;m bead enabled sufficient indentation of the sample to probe its stiffness while preserving its integrity. A bead with a smaller diameter can be used, after several optimizations, for gaining higher resolution in the stiffness maps but might lead to further overestimation of the Young&#39;s modulus values (for more details, see Crichton et al.65). Using the specified cantilever-bead ensemble, we were able to characterize sample stiffness in a broad range, from tens of units of Pa to ~100 kPa.
Sneddon&#39;s model was used to derive Young&#39;s modulus from force curves, as it allows analysis of deep indentations with colloidal probes62. Sneddon&#39;s model, unlike Hertz&#39;s model, does not suffer from the constraint that the contact radius must be much smaller than the sphere radius. It further assumes that the sample thickness is several times greater than the indentation depth30,66. In the present study, the indentation was ~2 &#181;m with a bead size of 5.7 &#181;m and a sample thickness of 30 &#181;m in collagen-rich areas; thus, Sneddon&#39;s model was appropriate. Other models63 considering the adhesion force between tip and substrate can be used for different types of tissues.
Analysis in AtomicJ implements corrections for the finite thickness of the samples to minimize the contribution of a substrate while deriving Young&#39;s modulus62,67. In the analysis of the obtained force curves, we used a single Poisson&#39;s ratio of 0.45, which has been previously recommended for soft tissue organs24. This approximation does not have a significant effect on calculated values of Young&#39;s modulus, as the change in the value of Poisson&#39;s ratio from 0.4 to 0.5 results only in a 0.893x decrease in Young&#39;s modulus values calculated according to the Sneddon&#39;s equation. Given the multi-fold differences in Young&#39;s modulus between the different durations of CCl4 treatments, the errors produced by approximating Poisson&#39;s ratio are only marginal.
We used withdraw curves to calculate stiffness values, as we were interested in the elastic response of the tissue to the load provided by the cantilever rather than in the plastic response to indentation68. Due to the viscoelastic response of the soft tissue, fitting withdraw curves might overestimate Young&#39;s modulus, which should be kept in mind. Furthermore, we have observed that data analysis with approach curves yields similar trends in stiffness values between fibrotic and control areas, though the absolute values are correspondingly lower (data not shown).
While optimizing the protocol, we identified several steps critical for the reproducibility of the measurements. First, it is important to ensure that the bead is approximately in the center of the translucent tip while attaching to the cantilever. This prevents possible mechanical imbalance during indentation. Second, during liver fixation with PFA, it is necessary to strictly follow the time limits for thawing and fixation. Changing the timing of this step might severely affect the mechanical properties of tissue sections. Third, the cantilever must be repeatedly calibrated with continuous monitoring and input of concurrent temperature values to avoid any artifacts occurring in stiffness values due to temperature fluctuations. Last, a single liver section should not be measured for longer than 3 h from preparation, as overlaid PBS may evaporate over longer periods. Readers can refer to the troubleshooting table (Table 3) for solving problems encountered during the AFM measurement, also discussed in length in Norman et al.46.
Table 3: Troubleshooting guide. <a href="https://www.jove.com/files/ftp_upload/63974/Table 3v2.xlsx" target="_blank">Please click here to download this Table.</a>
The presented protocol allows for reproducible AFM probing of liver tissue. It has the potential to reveal information on the development and eventual regression of fibrotic liver disease on a microscopic level and can contribute to the development of therapies targeting fibrotic scar regions formed during the progression of chronic liver disease.

The liver is a vital organ for maintaining homeostasis in organisms1,2. Chronic liver diseases account for ~2 million deaths worldwide annually3. They originate most commonly as viral infections, autoimmune disorders, metabolic syndromes, or alcohol abuse-related diseases and are accompanied by progressive liver fibrosis. Liver injury elicits an inflammatory response, which leads to the activation of cells depositing extracellular matrix (ECM) in a wound-healing response. However, in the presence of a chronic insult, excess ECM forms unresolved scar tissue within the liver, leading to the development of liver fibrosis, cirrhosis, liver carcinoma, and, ultimately, to liver failure4.
Hepatocyte injury immediately results in increased liver stiffness5,6. This directly affects hepatocyte function, activates hepatic stellate cells (HSCs) and portal fibroblasts, and results in their transdifferentiation to collagen-depositing myofibroblasts7,8. The deposition of fibrous ECM further increases liver stiffness, creating a self-amplifying feedback loop of liver stiffening and matrix-producing cell activation.
Liver stiffness has, thus, become an important parameter in liver disease prognosis. The change in biomechanical tissue properties can be detected earlier than fibrosis can be diagnosed by histological analysis. Therefore, various techniques for liver stiffness measurement have been developed in both research and clinical applications. In clinical settings, transient elastography (TE)9,10,11,12,13 and magnetic resonance elastography (MRE)14,15,16,17,18 have been employed to non-invasively diagnose early stages of liver damage by examining gross liver stiffness19.
In TE, ultrasound waves of mild amplitude and low frequency (50 Hz) are propagated through the liver, and their velocity is measured, which is then used to calculate tissue elastic modulus13. However, this technique is not useful for patients with ascites, obesity, or lower intercostal spaces due to improper transmission of the ultrasound waves through the tissues surrounding the liver9.
MRE is based on magnetic resonance imaging modality and uses 20-200 Hz mechanical shear waves to target the liver. A specific magnetic resonance imaging sequence is then used to trace the waves inside the tissue and to calculate the tissue stiffness16. Stiffness values reported with both TE and MRE techniques correlate well with the degree of liver fibrosis obtained from biopsies of human liver samples ranked using histological METAVIR scores20 (Table 1). TE and MRE have also been adapted for the measurement of liver stiffness in rodent models for research purposes21,22,23. However, as both methods derive the stiffness values from the tissue&#39;s response to the propagating shear waves, the values obtained might not reflect the absolute mechanical stiffness of the tissue.
For a direct mechanical characterization of rodent livers, Barnes et al. developed a model-gel-tissue assay (MGT assay) involving the embedding of liver tissue in polyacrylamide gel24. This gel is compressed by a pulsed uniform force from which Young&#39;s modulus can be calculated. The MGT assay shows a good correlation with an indentation assay adapted for both normal and fibrotic livers24 (Table 1).
Table 1: Liver stiffness values at the bulk level. TE and MRE compared to direct ex vivo mechanical measurements of liver elastic moduli using indentation and MGT assays for livers from different sources. The relation between E and G is given by E = 2G (1 + v), where v is the Poisson&#39;s ratio of the sample; F0 to F4 represent the fibrosis score in the METAVIR scoring system, with F0 denoting low or no fibrosis and F4 cirrhotic livers. Abbreviations: TE = transient elastography; MRE = magnetic resonance elastography; MGT = model-gel-tissue; E = elastic (Young&#39;s) modulus; G = shear modulus. <a href="https://www.jove.com/files/ftp_upload/63974/Table 1v2.xlsx" target="_blank">Please click here to download this Table.</a>
One of the major drawbacks of generic liver stiffness measurements is that they do not provide cellular-level resolution of stiffness heterogeneity in the liver. During the progression of fibrosis, collagen-rich areas show higher rigidity compared to the surrounding parenchyma25,26. This stiffness gradient locally influences the resident cells and plays an important role in driving HSC heterogeneity27. Thus, changes in local mechanical properties during liver disease development need to be characterized on a microscopic level to better understand fibrosis progression.
AFM allows the mechanical properties of tissue to be measured with high resolution and high force sensitivity. AFM uses the tip of a cantilever to indent the surface of a sample with forces as low as several piconewtons, inducing a deformation at a microscopic or nanoscopic level based on the geometry and size of the tip employed. The force response of the sample to the applied strain is then measured as the deflection in the cantilever28. Force-displacement curves are collected from the approach and retraction of the cantilever, which can be fitted with appropriate contact mechanics models to evaluate the local stiffness of the sample29.
In addition to measuring the stiffness of a given area, AFM can also provide topographic information about specific features in the sample, such as the structure of collagen fibres30,31,32. Multiple studies have described the application of AFM to measure the stiffness of various healthy and diseased tissues, such as skin32,33, lung34,35, brain36, mammary37,38,39, cartilage40, or heart41,42,43,44 from both patient and mouse model samples. Furthermore, AFM has also been used in vitro to determine the stiffness of cells and extracellular protein scaffolds45,46,47.
The measurement of the mechanical properties of biological samples using AFM is nontrivial due to their softness and fragility. Thus, various studies have standardized different conditions and settings, which yield widely fluctuating values of Young&#39;s moduli (reviewed by Mckee et al.48). Similar to other soft tissues, liver Young&#39;s modulus values at different grades of liver fibrosis also show extensive variation (Table 2). The differences in Young&#39;s modulus values arise from differences in the mode of AFM operation, cantilever tip, sample preparation method, sample thickness, indentation depth and forces, liver tissue environment during measurement, and analysis method (Table 2).
Table 2: Liver stiffness values at the cellular level. Liver stiffness values obtained using AFM describe the mechanical properties of the liver at the cellular level. Abbreviations: AFM = atomic force microscopy; E = elastic (Young&#39;s) modulus; PFA = paraformaldehyde; PBS = phosphate-buffered saline. <a href="https://www.jove.com/files/ftp_upload/63974/Table 2v2.xlsx" target="_blank">Please click here to download this Table.</a>
This paper describes a protocol for the reproducible measurement of Young&#39;s moduli of collagen-rich fibrotic areas in liver tissue by AFM with a precise localization provided by the use of polarization microscopy. We administered carbon tetrachloride (CCl4) to induce collagen deposition in a centrilobular fashion49 in a mouse model, reliably mimicking crucial aspects of human liver fibrosis50. Polarized microscopic images enable the visualization of collagen in the liver due to the birefringence of collagen fibers51, which allows accurate positioning of the cantilever tip over the desired area of interest within the hepatic lobule52.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
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			<td >AFM head</td>
			<td >Bruker</td>
			<td >JPK nanowizard 3</td>
			<td ></td>
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		<tr >
			<td >Cameras</td>
			<td >Andor</td>
			<td >Zyla 5.5 USB (sCMOS, water cooled)</td>
			<td></td>
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		<tr >
			<td ></td>
			<td >The Imagingsource&#160;</td>
			<td >S/N:12310015</td>
			<td></td>
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		<tr >
			<td >Cantilever</td>
			<td >SD-qp-BioT-TL-10, Nanosensors</td>
			<td>S/N:73750F05</td>
			<td></td>
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		<tr >
			<td >Cryotome</td>
			<td >Leica&#160;</td>
			<td>CM1950</td>
			<td></td>
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		<tr >
			<td >Epoxy resin glue (Long working time )</td>
			<td >Bison epoxy universal</td>
			<td></td>
			<td></td>
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		<tr >
			<td >Melamine beads; diameter, 5.7 um</td>
			<td >Microparticles, GmbH</td>
			<td>MF-R-5.7</td>
			<td></td>
		</tr>
		<tr >
			<td >Microscope</td>
			<td >Olympus</td>
			<td>&#160;IX81</td>
			<td></td>
		</tr>
		<tr >
			<td >Hydrophobic&#160; slide marker</td>
			<td >SuperHT</td>
			<td>PAP PEN</td>
			<td></td>
		</tr>
		<tr >
			<td >Software</td>
			<td >JPK nanowizard&#160; v6.1.151</td>
			<td></td>
			<td></td>
		</tr>
		<tr >
			<td ></td>
			<td >AtomicJ v2.3.1</td>
			<td></td>
			<td></td>
		</tr>
		<tr >
			<td >Superfrost slides</td>
			<td >Thermoscientific</td>
			<td>ref no. J1800AMNZ</td>
			<td></td>
		</tr>
		<tr >
			<td >System</td>
			<td >Ubuntu 14.04.5 LTS</td>
			<td></td>
			<td></td>
		</tr>
		<tr >
			<td >Vibration isolation control unit</td>
			<td >Tablestable</td>
			<td>AVI-200-S</td>
			<td></td>
		</tr>
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</table>

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.

Mildly fixed, 30 &#181;m thick liver sections obtained from control mice and from mice with mild or advanced fibrosis (induced by injection of CCl4 for 3 weeks or 6 weeks, respectively49) were probed with AFM as described in this protocol. Collagen fibers close to central veins were selected for the measurement of stiffness maps. Areas close to the central veins, which correspond to the areas where collagen fibers in CCl4-treated animals usually form, were analyzed in control livers (Figure 4A). The distribution of Young&#39;s moduli was reproducible across different control livers and collagen-rich areas within a single liver section (Figure 4B: left violin plot).
In CCl4-treated animals, stiffness maps corresponding to the pericentral areas of collagen deposits showed significantly higher values of Young&#39;s moduli compared to equivalent areas in control mice (Figure 4B: 1.9 kPa vs. 2.6 kPa median Young&#39;s modulus values for control vs. 3 week CCl4-treated mouse; p = 0.07; 1.9 kPa vs. 5.1 kPa median Young&#39;s modulus values for control vs. 6 weeks CCl4-treated mouse; p = 0.02). Moreover, there was a significant increase in the values of Young&#39;s moduli with longer CCl4 treatment (Figure 4B; 2.6 kPa vs. 5.1 kPa median Young&#39;s modulus values for 3 week vs. 6 week CCl4-treated mouse; p = 0.04). This shows a gradual stiffening of collagen deposits with fibrosis progression and that AFM measurements reflect the fibrogenesis.
To evaluate the effect of prolonged storage of OCT-embedded liver sections on the mechanical properties of collagen fibers, we measured the stiffness of collagen fibers in sections of CCl4-treated mice, which were stored at &#8722;80 &#176;C for 2 weeks or 3 months on the slide after cutting (Figure 5). AFM measurements showed significantly lower values of Young&#39;s moduli in collagen-rich areas for sections stored for 3 months compared to those obtained from sections measured within 2 weeks of sample sectioning (Figure 5; 4.7 kPa vs. 3.6 kPa median Young&#39;s modulus values for 2 weeks vs. 3 months storage; p &#60; 0.001). Thus, it is important to measure the mechanical properties of the liver tissue shortly after the sections are prepared from the OCT-embedded liver lobes.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63974/63974fig04.jpg" /> 
Figure 4: AFM measurements reveal progressive stiffening of collagen-rich areas correlating with prolonged CCl4 treatment.&#160;(A)&#160;Liver sections from control mice and mice treated with CCl4 for 3 weeks or 6 weeks were used to measure the mechanical properties of collagen-rich areas. The boxed areas of liver sections shown in the polarized microscopy images (left) are collagen-rich scan areas (or corresponding regions in the control liver) selected for AFM measurements (30 &#181;m&#160;x 100 &#181;m, 10 pixels x 36 pixels). The Young&#39;s modulus maps with color scales corresponding to these boxed areas are shown on the right, including histograms of Young&#39;s modulus values from these maps; inset scatter plots show values &#62;10 kPa for each condition. The stiffening of the liver is visualized as a gradual rightward shift in the histogram distribution and a higher frequency of points in the inset scatter plot. Scale bar = 20 &#181;m. (B)&#160;Violin plots show the distribution of elastic moduli from three areas measured for each condition (left) and summarized elastic moduli values from all three maps (right). Violin plots show the median (red line), 25th percentile&#160;and 75th percentile (black lines); dots represent the mean values of individual maps from areas 1-3. The presented p-values were calculated using a Student&#39;s t-test performed on means. Abbreviations: AFM = atomic force microscopy; CCl4 = carbon tetrachloride. <a href="https://www.jove.com/files/ftp_upload/63974/63974fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63974/63974fig05.jpg" /> 
Figure 5: Extended storage of liver sections leads to a decrease in stiffness of collagen-rich areas.&#160;Liver sections (prepared from mice treated with CCl4 for 2 weeks) stored at &#8722;80 &#176;C for 2 weeks or 3 months were used to measure Young&#39;s modulus. Polarized microscopy images (left) with boxes indicating the collagen-rich areas used for AFM measurement (30 &#181;m&#160;x 100 &#181;m, 10 pixels x 36 pixels). Corresponding Young&#39;s modulus maps with color scales (right). Histograms show Young&#39;s modulus values collected from 4-6 areas in each sample; inset scatter graphs show values &#62;10 kPa for each condition. Scale bar = 20 &#181;m.&#160;Abbreviations: AFM = atomic force microscopy; CCl4 = carbon tetrachloride. <a href="https://www.jove.com/files/ftp_upload/63974/63974fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure S1: Method for modifying the cantilever with a melamine resin micro bead. (A)&#160;Drawn schematic illustrates the attachment of a spherical bead to the tip of the cantilever. For a step-wise description, see Section 2, Part 1, Attachment of a 5.7 &#181;m bead to AFM cantilever tip. (B)&#160;Microscopy image of a spherical 5.7 &#181;m bead attached to the cantilever tip shown from top (left) and lateral view (right). Scale bars = 20 &#181;m. Abbreviations: AFM = atomic force microscopy; RT = room temperature. <a href="https://www.jove.com/files/ftp_upload/63974/supplementary fig 1.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Data analysis in AtomicJ. (A) The sequence of steps to be followed for opening stiffness maps in AtomicJ. A single left-click on the process force curves and maps (x) opens the processing assistant. Files can be loaded to the processing assistant by clicking on add&#160;(y) and selecting the required files. Click on next&#160;(z) to proceed to the next step. (B)&#160;Parameters for curve fitting, appropriate contact mechanics model, and AFM settings used during measurement. Steps 1-11&#160;refer to corresponding sub-points detailed in protocol step 3, Section 5, Data analysis. <a href="https://www.jove.com/files/ftp_upload/63974/supplementary_fig_2.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S3: Outline of analyzed data in AtomicJ. The preview of analyzed data shows stiffness maps (upper left window), force curves (upper right window), and raw data (lower window). Abbreviation: AFM = atomic force microscopy. <a href="https://www.jove.com/files/ftp_upload/63974/supplementary fig 3.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy at JoVE.com

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

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 ...

measurement-liver-stiffness-using-atomic-force-microscopy-coupled

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3.6K Views.  Institute of Molecular Genetics of the Czech Academy of Sciences.  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.

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

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 ...

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.

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 ...

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 ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

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 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 ...