Name: measurement of liver stiffness using atomic force microscopy coupled with polarization microscopy
Uploaded: 2022-07-20T14:00:00
Duration: 10 min 10 s
Description: Watch this Scientific Journal Video about Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy at JoVE.com

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.

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			<td >Zyla 5.5 USB (sCMOS, water cooled)</td>
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			<td >The Imagingsource&#160;</td>
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			<td >Cantilever</td>
			<td >SD-qp-BioT-TL-10, Nanosensors</td>
			<td>S/N:73750F05</td>
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			<td >Cryotome</td>
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			<td>CM1950</td>
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			<td >Epoxy resin glue (Long working time )</td>
			<td >Bison epoxy universal</td>
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			<td >Melamine beads; diameter, 5.7 um</td>
			<td >Microparticles, GmbH</td>
			<td>MF-R-5.7</td>
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			<td >Olympus</td>
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			<td >Hydrophobic&#160; slide marker</td>
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			<td >JPK nanowizard&#160; v6.1.151</td>
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			<td >AtomicJ v2.3.1</td>
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			<td >Superfrost slides</td>
			<td >Thermoscientific</td>
			<td>ref no. J1800AMNZ</td>
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			<td >System</td>
			<td >Ubuntu 14.04.5 LTS</td>
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			<td >Vibration isolation control unit</td>
			<td >Tablestable</td>
			<td>AVI-200-S</td>
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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 ...

Our protocol provides a distinction between collagen rich and collagen lacking areas in the liver sections. It can thus be used to study the processes of fibrogenesis in the greater details. The main advantage of the technique lies in the use of polarizing microscopy to locate collagen rich areas in the liver sections.Our protocols can provide great insight into the development of liver fibrosis, and it can also be adopted for other soft tissues such as lungs after certain optimizations. To begin, remove the frozen liver sections from minus 80 degrees Celsius, and thaw them at room temperature for two minutes. Fix the sections with ice cold 4%paraformaldehyde in PBS for 10 minutes at four degrees Celsius, followed by washing five times with PBS.After the last wash, wipe the excess PBS around the liver section using tissue, and mark a boundary of approximately two by four centimeters around the liver section with a hydrophobic marker pen. Cover the sample with PBS, then load the sample onto the AFM stage, and cover it with the AFM head. In the atomic force microscopy or AFM software, go to the setup tab and mark a tick on autosave to automatically save all files.Then specify the file name and directory for saving the measurement files by going to the setup tab and clicking on saving settings. Approach the tissue surface with the AFM cantilever by switching on the laser and clicking on the approach key. After the approach is complete, retract the cantilever tip by clicking on the retraction key once, which retracts the cantilever to the top end of the piezo range.If required, realign the detector using knobs on the AFM head. Turn off the laser. The tip is now in the focus of the objective.Next, overlay the optical field of the microscope with AFM measurement maps by clicking on the accessories tab and selecting direct overlay optical calibration. In the succeeding window, click next to take a series of images of the cantilever scanning a specific area. Click next again to go to the next window.In the first image, click on the center of the tip of the cantilever manually to depict the tip position in the software. The circle depicting the tip position can be manipulated in size by indicating its radius to increase accuracy. Click calibrate to detect the cantilever tip position in all images automatically.Confirm the accuracy of tip detection by going through the images. Click next, and then finish to finish the optical overlay. Next, move the stage to position a collagen rich area inside the green box, visible in the data viewer tab, using the polarized image.Select the collagen rich area by defining the area under the grid tab, and then making a rectangle on the data viewer tab using a long press on the left mouse button inside the specified green box. Click on confirm new scan region to set the selected area as an area of measurement. Set the parameters of the feedback loop.Set I gain at 50 hertz and P gain at 0.001. Then set the set point at one nano Newton. Select the relative set point value according to the studied materials mechanical properties and the cantilever's stiffness.Specify a just baseline to fit the force curves properly. Then select the length of the cantilever movement in the Z axis according to the surface topography of the sample. Set Z movement to constant duration.Set the extend time to one second with extension and retraction delays as zero. Set sample rate at 5, 000 hertz. Mark a tick in front of Z closed loop to automatically adjust the distance between the sample and the cantilever tip during measurements.Then disengage the motorized stage control by deselecting engage button. Switch on the laser and approach the sample with the cantilever. Click on start scan to collect force distance curves.Analyze the acquired data using the open source software AtomicJ. Load the force curves into the program by clicking on the process force curves and maps icon. Then in the processing assistant, add the maps to be analyzed by clicking on the add button.After the maps are loaded, click on next. Specify the processing settings in the next window. To estimate the contact point between the sample and the cantilever automatically using a set of fitting curve parameters, use the automatic estimation of the contact point.Determine the contact point between the cantilever and the sample by the classical focused grid method. Select the estimation method as the model independent method to yield the best determination of the contact point based on the quality of the measured force curves, which needs to be empirically determined during the optimization of data analysis. Fit the force indentation curve using a classical model as classical L two for model fit.Set the fit of the model to the withdrawal curve. Set Poisson's ratio as 0.45 as recommended for soft tissues such as the liver. Set the fit of the curve using a baseline degree of three and an in-contact degree of one.Change the degree of polynomial fit based on the scale of deviations of the curves from the model. Select the model used to fit the withdrawal curves as the Sneddon model. Fill in the radius of the spherical tip to 2.9 micrometers.Load the spring constant and invOLS from the data files by enabling read-in, then click on finish. After data analysis, go through the force curves. Exclude the force curves where the cantilever approached the surface of the liver section incorrectly.These curves have high noise and aberrant shapes. After this, the data can be copied or exported. In this study, mildly fixed liver sections obtained from the control mice and mice with mild and advanced fibrosis induced by the injection of carbon tetrachloride were probed with AFM.Areas close to the central veins corresponding to the areas where collagen fibers in carbon tetrachloride mice usually form were analyzed in control livers. The distribution of Young's moduli was reproducible across different regions in control livers and collagen rich areas within a single liver section. In carbon tetrachloride treated mice, stiffness maps corresponding to the pericentral areas of collagen deposits showed significantly higher values of Young's moduli compared to equivalent areas in control mice.Moreover, there was a significant increase in the values of Young's moduli with longer treatment. The effect of prolonged storage of liver sections on the mechanical properties of collagen fibers was also assessed. AFM measurements showed significantly lower values of Young's moduli in collagen rich areas for sections stored for three months, compared to those obtained within two weeks.The time of fixation of liver section greatly determines the mechanical properties of the liver section. We recommend timed intervals stated in the protocol must be strictly followed. After certain optimizations in the protocol, the given method can be used for any soft tissues showing significant collagen deposition, which is detectable by polarizing microscopy.The given technique provides the researchers with a unified and convenient protocol to study mechanism of liver across healthy and disease.

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In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

In vivo Quantification of G Protein Coupled Receptor Interactions using Spectrally Resolved Two-photon Microscopy

The study of protein interactions in living cells is an important area of research because the information accumulated both benefits industrial ...

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 ...

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 ...

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 ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

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

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 ...

Detection of Ligand-activated G Protein-coupled Receptor Internalization by Confocal Microscopy

Confocal laser scanning microscopy (CLSM) is an optical imaging technique for high-contrast imaging. It is a powerful approach to visualize fluorescent ...