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><tbody><tr><td>AFM head</td><td>Bruker</td><td>JPK nanowizard 3</td><td></td></tr><tr><td>Cameras</td><td>Andor</td><td>Zyla 5.5 USB (sCMOS, water cooled)</td><td></td></tr><tr><td></td><td>The Imagingsource&amp;nbsp;</td><td>S/N:12310015</td><td></td></tr><tr><td>Cantilever</td><td>SD-qp-BioT-TL-10, Nanosensors</td><td>S/N:73750F05</td><td></td></tr><tr><td>Cryotome</td><td>Leica&amp;nbsp;</td><td>CM1950</td><td></td></tr><tr><td>Epoxy resin glue (Long working time )</td><td>Bison epoxy universal</td><td></td><td></td></tr><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>&amp;nbsp;IX81</td><td></td></tr><tr><td>Hydrophobic&amp;nbsp; slide marker</td><td>SuperHT</td><td>PAP PEN</td><td></td></tr><tr><td>Software</td><td>JPK nanowizard&amp;nbsp; 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></tbody></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

Research

JoVE Journal

Biology

3.7K 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

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

Covalent Attachment of Single Molecules for AFM-based Force Spectroscopy

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and surfaces. For a true single molecule experiment, covalent attachment of the probe molecule is essential because only then can hundreds of force-extension traces with one and the same single molecule be obtained. Many traces are in turn necessary to prove that a single molecule alone is probed. Additionally, passivation is crucial for preventing unwanted interactions between the single probe molecule and the AFM cantilever tip as well as between the AFM cantilever tip and the underlying surface. The functionalization protocol presented here is reliable and can easily be applied to a variety of polymers. Characteristic single molecule events (i.e., stretches and plateaus) are detected in the force-extension traces. From these events, physical parameters such as stretching force, desorption force and desorption length can be obtained. This is particularly important for the precise investigation of stimuli-responsive systems at the single molecule level. As exemplary systems poly(ethylene glycol) (PEG), poly(N-isopropylacrylamide) (PNiPAM) and polystyrene (PS) are stretched and desorbed from SiOx (for PEG and PNiPAM) and from hydrophobic self-assembled monolayer surfaces (for PS) in aqueous environment.

Covalent attachment of probe molecules to atomic force microscopy (AFM) cantilever tips is an essential technique for the investigation of their physical properties. This allows us to determine the stretching force, desorption force and length of polymers via AFM-based single molecule force spectroscopy with high reproducibility.

Atomic force microscopy (AFM)-based single molecule force spectroscopy is an ideal tool for investigating the interactions between a single polymer and ...

Chemistry

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 ...

Engineering

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 ...

Application of Atomic Force Microscopy to Detect Early Osteoarthritis

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in elasticity can propagate or trigger the onset of major diseases like cancer or osteoarthritis (OA). Atomic force microscopy (AFM) has emerged as a strong tool to qualitatively and quantitatively characterize the biomechanical properties of specific biological target structures on a microscopic scale, measuring forces in a range from as small as the piconewton to the micronewton. Biomechanical properties are of special importance in musculoskeletal tissues, which are subjected to high levels of strain. OA as a degenerative disease of the cartilage results in the disruption of the pericellular matrix (PCM) and the spatial rearrangement of the chondrocytes embedded in their extracellular matrix (ECM). Disruption in PCM and ECM has been associated with changes in the biomechanical properties of cartilage. In the present study we used AFM to quantify these changes in relation to the specific spatial pattern changes of the chondrocytes. With each pattern change, significant changes in elasticity were observed for both the PCM and ECM. Measuring the local elasticity thus allows for drawing direct conclusions about the degree of local tissue degeneration in OA.

We present a method to investigate early osteoarthritic changes at the cellular level in articular cartilage by using atomic force microscopy (AFM).

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in ...

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 ...

Quantitative Analysis of Liver Stiffness Distribution

A Three-Dimensional Digital Model for Early Diagnosis of Hepatic Fibrosis Based on Magnetic Resonance Elastography

Hepatic fibrosis is an early stage of liver cirrhosis, and there are no better non-invasive and convenient methods for the detection and evaluation of the disease. Despite the good progress made with the liver stiffness map (LSM) based on magnetic resonance elastography (MRE), there are still some limitations that need to be overcome, including manual focus determination, manual selection of regions of interest (ROIs), and discontinuous LSM data without structural information, which makes it impossible to evaluate the liver as a whole. In this study, we propose a novel three-dimensional (3D) digital model for the early diagnosis of hepatic fibrosis based on MRE.
MRE is a non-invasive imaging technique that employs magnetic resonance imaging (MRI) to measure the liver stiffness at the scanning site through human-computer interaction. Studies have indicated a significant positive correlation between the LSM obtained through MRE and the degree of hepatic fibrosis. However, for clinical purposes, a comprehensive and precise quantification of the degree of hepatic fibrosis is necessary. To address this, the concept of Liver Stiffness Distribution (LSD) was proposed in this study, which refers to the 3D stiffness volume of each liver voxel obtained by the alignment of 3D liver tissue images and MRE indicators. This provides a more effective clinical tool for the diagnosis and treatment of hepatic fibrosis.

Author Spotlight: Advancing Hepatic Fibrosis Diagnosis Using Magnetic Resonance Elastography and AI 

The objective of this study was to develop a novel three-dimensional digital model for the early diagnosis of hepatic fibrosis, which includes the stiffness of each voxel in the patient's liver and can thus, be used to calculate the distribution ratio of the patient's liver at different fibrosis stages.

Hepatic fibrosis is an early stage of liver cirrhosis, and there are no better non-invasive and convenient methods for the detection and evaluation of the ...

Medicine

Application of Ultrasound and Shear Wave Elastography Imaging in a Rat Model of NAFLD/NASH

Nonalcoholic Steatohepatitis (NASH) is a condition within the spectrum of Non-Alcoholic Fatty Liver Disease (NAFLD), which is characterized by liver fat accumulation (steatosis) and inflammation leading to fibrosis. Preclinical models closely recapitulating human NASH/NAFLD are essential in drug development. While liver biopsy is currently the gold standard for measuring NAFLD/NASH progression and diagnosis in the clinic, in the preclinical space, either collection of whole liver samples at multiple timepoints during a study or biopsy of liver is needed for histological analysis to assess the disease stage.
Conducting a liver biopsy mid-study is an invasive and labor-intensive procedure, and collecting liver samples to assess disease level increases the number of research animals needed for a study. Thus, there is a need for a reliable, translatable, non-invasive imaging biomarker to detect NASH/NAFLD in these preclinical models. Non-invasive ultrasound-based B-mode images and Shear Wave Elastography (SWE) can be used to measure steatosis as well as liver fibrosis. To assess the utility of SWE in preclinical rodent models of NASH, animals were placed on a pro-NASH diet&#160;and underwent non-invasive ultrasound B-mode and shear wave elastography imaging to measure hepatorenal (HR) index and liver elasticity, measuring progression of both liver fat accumulation and tissue stiffness, respectively, at multiple time points over the course of a given NAFLD/NASH study.
The HR index and elasticity numbers were compared to histological markers of steatosis and fibrosis. The results showed strong correlation between the HR index and percentage of Oil Red O (ORO) staining, as well as between elasticity and Picro-Sirius Red (PSR) staining of livers. The strong correlation between classic ex vivo methods and in vivo imaging results provides evidence that shear wave elastography/ultrasound-based imaging can be used to assess disease phenotype and progression in a preclinical model of NAFLD/NASH.

This protocol describes the use of an enhanced ultrasound technique to non-invasively observe and quantify liver tissue changes in rodent models of nonalcoholic fatty liver disease.

Nonalcoholic Steatohepatitis (NASH) is a condition within the spectrum of Non-Alcoholic Fatty Liver Disease (NAFLD), which is characterized by liver fat ...

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 ...

Ultrasound II: Endoscopic Ultrasound and FibroScan

Endoscopic Ultrasound (EUS) and FibroScan are valuable diagnostic tools in gastroenterology and hepatology, each with specific applications and techniques.
Endoscopic Ultrasound (EUS):
<ol style="list-style-type: circle;">
	<li>Technological Integration: EUS is a sophisticated integration of optical endoscopy and ultrasonography. The endoscope visualizes the mucosal surface directly, while the ultrasound component penetrates deeper to visualize organs and structures beyond the gastrointestinal wall.</li>
	<li>Image Formation: The ultrasound component emits high-frequency sound waves ranging from 5 to 20 MHz. These waves reflect differently from various types of tissues, creating detailed cross-sectional images of the internal structures.</li>
	<li>Layered Visualization: EUS is particularly good at visualizing the five distinct layers of the gastrointestinal wall, which is crucial for staging gastrointestinal cancers, especially esophageal, gastric, and rectal cancers.</li>
	<li>Fine Needle Aspiration (FNA): EUS can be combined with FNA, in which a thin needle is passed via the endoscope to collect tissue samples from lymph nodes or masses. This technique is invaluable for staging cancer, diagnosing unexplained masses, guiding treatment decisions, and tailoring therapeutic strategies.</li>
	<li>Therapeutic Applications: Beyond diagnostics, EUS can be used for therapeutic procedures like draining cysts or pseudocysts, performing nerve blocks for chronic pancreatitis pain management, and delivering medication directly into tumors.</li>
</ol>
FibroScan (Transient Elastography):
<ol style="list-style-type: circle;">
	<li>Principle of Operation: FibroScan measures liver stiffness by emitting mechanical vibrations of mild amplitude and low frequency (approximately 50 Hz). It measures the velocity of the shear wave (elastic wave) generated by these vibrations.</li>
	<li>Elasticity Measurement: The speed of the shear wave correlates with liver tissue stiffness. The stiffer the tissue (as in fibrosis), the faster the shear wave travels. This speed is converted into a liver stiffness measurement (LSM), usually expressed in kilopascals (kPa).</li>
	<li>CAP (Controlled Attenuation Parameter): Some FibroScan models also feature CAP, which estimates the liver&#39;s degree of steatosis (fat content). It provides a quantitative assessment that is particularly useful for diagnosing and monitoring non-alcoholic fatty liver disease (NAFLD).</li>
	<li>Diagnostic Utility: FibroScan provides quantitative metrics critical for evaluating the severity of liver fibrosis, especially in chronic liver diseases like hepatitis B and C. It helps monitor disease progression and assess the effectiveness of antifibrotic therapies.</li>
	<li>Advantages Over Liver Biopsy: FibroScan is noninvasive, has a minimal risk of complications compared to liver biopsy, and is more patient-friendly. It can also sample a larger volume of liver tissue, potentially providing a more representative assessment of liver stiffness than a biopsy. However, it may be less accurate in cases of severe obesity or ascites.</li>
</ol>
Both EUS and FibroScan represent significant advancements in gastroenterology and hepatology. EUS offers a unique combination of endoscopic visualization and deep tissue ultrasound imaging, making it an essential tool for diagnosing and staging gastrointestinal diseases. Meanwhile, FibroScan provides a quick, non-invasive, and reliable method for assessing liver health, particularly in fibrosis and steatosis.

Endoscopic ultrasound is a minimally invasive diagnostic procedure that combines endoscopy and ultrasound to visualize the digestive tract and nearby ...

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

Summary

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

A Three-Dimensional Digital Model for Early Diagnosis of Hepatic Fibrosis Based on Magnetic Resonance Elastography

Application of Ultrasound and Shear Wave Elastography Imaging in a Rat Model of NAFLD/NASH

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

Ultrasound II: Endoscopic Ultrasound and FibroScan