We thank the orthopedic surgeons from the Department of Orthopaedic Surgery of the University Hospital of Tuebingen for providing the tissue samples.

Femoral condyles collected from patients undergoing total knee arthroplasty at the University Hospital of T&#252;bingen, Germany, were used. Only articular cartilage samples from patients with degenerative and posttraumatic joint pathologies were included in this study. Departmental, institutional, as well as local ethical committee approval were obtained before the commencement of the study (Project no.674/2016BO2). Written informed consent was received from all patients before participation.
NOTE: A flowchart of the experiment steps in their chronological order is given in Figure 1.
1. Tissue processing and generation of cartilage discs
<ol>
	<li>Tissue preparation
	<ol>
		<li>Following post-operative resection, place the cartilage samples in a container filled with Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) supplemented with 5% (v/v) penicillin-streptomycin. Make sure the samples are completely submerged in the medium. The duration between surgical resection and further processing of the cartilage should not exceed 24 h. Ensure that, throughout the entire processing, the samples are fully submerged in media to avoid sample drying.</li>
		<li>Cut the cartilage away from the bone using a scalpel.</li>
	</ol>
	</li>
	<li>Cartilage disc generation
	<ol>
		<li>Generate cartilage discs of 4 mm in diameter using a biopsy punch. 
		NOTE: It is important to select and resect the areas of the condyle where the cartilage layer thickness exceeds 1 mm. This might be problematic, especially around loadbearing zones, where the cartilage layer typically loses its thickness due to wear and tear processes or degeneration.</li>
		<li>Place the previously generated 4 mm cartilage discs on a custom-made cutting device and fix and hold the cartilage discs stable by means of a spatula. When placing the cartilage discs on the cutting device, care must be taken. Position the samples so that the cartilage&#39;s topmost layer (the superficial zone of the articular cartilage) does not face the blade</li>
		<li>Cut the cartilage discs with a razor blade. Disc-shaped cartilage samples of 4 mm x 1 mm are, thus, generated. To prevent sample drying, perform tissue cutting as quickly as possible.</li>
		<li>Collect each disc with the help of a spatula and place the generated cartilage discs into 1.5 mL tubes containing 1 mL of DMEM supplemented with 5% (v/v) penicillin-streptomycin. Place approximately 15 discs in one tube.</li>
	</ol>
	</li>
	<li>Cryotome sectioning of the cartilage discs (for perpendicular slices) 
	NOTE: This step is optional, and it can be employed if a side-view visualization of the cellular pattern distribution within the cartilage discs is desired. It can be used as a verification method as the distribution of cellular pattern is a 3D feature of articular cartilage26. Optical sectioning and 3D reconstructions of the entire cartilage discs using a confocal microscope can also be used, removing, thus, the need to section the samples as described in the protocol.
	<ol>
		<li>Cover the cartilage disc with water-soluble embedding medium and place it on its edge on the cryotome knob (with the surface of the disc perpendicular to the surface of the knob). In the cryotome device, the embedding medium freezes at low temperatures.</li>
		<li>Using a standard cryotome, section the tissue sideways at a 60 &#181;m thickness until the middle of the disc is reached (i.e., when cryosections reach a length of 4 mm) and collect the slices. By sectioning the disc explant perpendicularly, all zones of the cartilage (superficial, middle, and deep) can be visualized.</li>
		<li>Collect the sections on a glass slide and remove the water-soluble embedding medium by washing three times with phosphate-buffered saline (PBS).</li>
	</ol>
	</li>
</ol>
2. Cartilage disc sorting as a function of the cellular spatial pattern
<ol>
	<li>Staining of the disc-shaped cartilage samples
	<ol>
		<li>Place one cartilage disc (section 1.2) in each well of a 96-well plate and add 130 &#181;L of cell permeable fluorescence dye at a dilution of 1:1,000 to each well.</li>
		<li>Visually inspect the entire plate and make sure that only one disc is placed in each well. Incubate the plate for 30 min in the standard cell culture incubator at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Staining of the 60 &#181;m cartilage slices
	<ol>
		<li>Gently place the cartilage disc sections (section 1.3) on glass microscope slides with the help of forceps.</li>
		<li>Cover the cartilage sections with mounting medium containing nuclear DAPI counterstaining and gently place coverslips that are suitable for fluorescence microscopy.</li>
		<li>Seal the edges of each coverslip with regular transparent nail polish and allow to dry for 3 min.</li>
	</ol>
	</li>
	<li>Top-down and side-view cartilage sorting and imaging 
	NOTE: Each disc must be examined under a fluorescent microscope. The aim of this step is to sort the discs based on their predominant cellular pattern (single strings, double strings, small clusters, big clusters, or diffuse).
	<ol>
		<li>Place the 96-well plate on the plate holder of the fluorescence microscope.</li>
		<li>Select the appropriate fluorescence filter of either Em 495 nm/Ex 515 nm (for top-down imaging of the cartilage discs prepared in section 2.1.) or Em 358 nm/Ex 461 nm (for side-view imaging of the cartilage sections prepared in section 2.2) and the 10x objective. 
		NOTE: Using the 10x objective allows the whole circumference of the disc to be inspected, and samples with inhomogeneous or improper staining can be excluded. However, using only the top-down view may result in the perception of changes in cellular organization as a result of analysis of the deeper tissue layers made visible to top-down observation by superficial erosion. As an example, an ascending string following the collagen arcades could be perceived as a single cell or scattered cells (diffuse pattern)26. As a result, both sides of the discs must be inspected to ensure proper cellular pattern selection.</li>
		<li>Determine visually the cellular pattern displayed in each cartilage disc. It is unlikely that a disc will have only one type of cellular pattern. For the portion of the disc where the chondrocyte arrangement does not match the pattern of interest, only accept the samples if the undesired pattern is at the very periphery, where AFM measurements are not taking place (i.e., up to 0.5 mm from the disc border), and ensure that this does not exceed 10% of the total surface of the disc27,28.</li>
	</ol>
	</li>
	<li>Image acquisition of the entire cartilage discs
	<ol>
		<li>Select the 10x objective of the microscope and position it underneath the preselected well containing an individual cartilage disc. Focus on the disc to see the cellular pattern.</li>
		<li>Select the Navigator Function to get an overview of the entire well. Use the left mouse button and drag to navigate to a different stage position. With the mouse wheel, zoom in and out. 
		NOTE: At this point, a preview of the well with the entire sample can be seen by double-clicking on each area of interest sequentially.</li>
		<li>Select a square that encompasses the area of interest to be scanned; at this point, all single tiles that compose the mosaic will become visible.</li>
		<li>Adjust the exposure/light intensity so that the cells can be clearly visualized from the background. At this point, the picture&#39;s brightness/contrast has been adjusted for all the tiles and can no longer be customized individually for each tile. 
		NOTE: As the cells near the disc&#39;s edge often emit a higher fluorescence signal than the cells in the center, the exposure/light intensity settings must be adapted.
		<ol>
			<li>To evaluate if the exposure time is appropriate for a particular channel, examine the distribution of the signal in the histogram. By using the automatic exposure mechanism included in the microscope&#39;s image software, visualize all the cells residing within the disc.</li>
		</ol>
		</li>
		<li>Select the software&#39;s Focus Map Point option, and then select each individual tile by left-clicking in the center of it.</li>
		<li>Select the option Focus Map. A window is displayed with all the previously selected tiles. Double-click on a tile in the list to display and bring it into proper focus.</li>
		<li>Click Set Z to save the focal plan and proceed to the next tile. After adjusting the focal plan for each individual tile, begin image acquisition by pressing Start Scan.
		<ol>
			<li>If the scan is displaying darker horizontal and/or vertical bars, this may be due to improper and uneven illumination of the single frames. Resolve this by using the Linked Shading option incorporated in the software prior to the actual scan.</li>
		</ol>
		</li>
		<li>Save, export, and correctly annotate the images.</li>
	</ol>
	</li>
</ol>
3. Biomechanical approach of cartilage explants
<ol>
	<li>Sample preparation for AFM measurements
	<ol>
		<li>Fix each preselected cartilage disc containing a cellular pattern (section 2) in Petri dishes by means of biocompatible glue.Add sufficient sample glue on the top, bottom, left, and right sides of the disc.</li>
		<li>Cover the discs with 2.5 mL of Leibovitz&#39;s L-15 medium without L-glutamine. Add the Leibowitz medium gently onto the samples to avoid sample detachment from the surface due to waves created by the medium.</li>
	</ol>
	</li>
	<li>Loading of the samples into the AFM
	<ol>
		<li>Position the Petri dish in the AFM device&#39;s sample holder and turn on the Petri dish heater set to 37 &#176;C. Allow the tissue culture dish to reach the desired temperature. This is done to exclude possible artifacts caused by temperature variation.</li>
	</ol>
	</li>
	<li>AFM-cantilever calibration
	<ol>
		<li>Initialize the software setup as previously described by Danalache et al.29.</li>
		<li>Select a suitable glass block cantilever holder for liquid measurements and carefully place it on the AFM head. A locking mechanism secures the glass block in the AFM head. Ensure that the glass block&#39;s reflective surface is straight and parallel to the AFM holder.</li>
		<li>Place the cantilever on the surface of the glass block cantilever holder with care. The cantilever itself should rest on the polished optical plane, in the center of the glass block.</li>
		<li>Carefully place a silicone skirt (silicone membrane) on the base of the cantilever holder in order to prevent medium condensation in the AFM head.</li>
		<li>Lower the cantilever in 100 &#181;m steps using the stepper motor function until it is completely submerged in the medium.</li>
		<li>Run a scanner approach with the approach parameters described by Danalache et al.29. Retract the cantilever by 100 &#181;m once the bottom of the Petri dish is reached.</li>
		<li>Calibrate the cantilever using the exact steps and run the parameters described by Danalache et al.29. At the end of the calibration, the vertical deflection is saved and displayed in newton (N) units of force rather than volts (V)-the unit of the original registration by the photodiode detector. In the experiments here, a set point of 4.47 nN resulted after calibration.</li>
		<li>Using the stepper motor function, retract the cantilever to 1,000 &#181;m.</li>
	</ol>
	</li>
	<li>Identifying the desired cartilage measurement site under the AFM 
	NOTE: Due to the 1 mm thickness of the cartilage discs, the cantilever is not visible in the field of view while navigating over the sample.
	<ol>
		<li>Use the CCD camera of the microscope to identify the cantilever. The AFM cantilever should be positioned in a sample-free area of the Petri dish.</li>
		<li>Start a scanner approach with the cantilever on a clean, sample-free area of the Petri dish, using the same parameters described by Danalache et al.29.</li>
		<li>Further retract the cantilever 1.5 mm away from the bottom of the plate with the stepper motor control. This step is crucial in order to avoid a direct collision between the cantilever and the sample.</li>
		<li>Switch from brightfield to fluorescence view and visually identify the top of the disc.</li>
		<li>Move the AFM sample holder exactly 2 mm toward the middle of the disc. This point is considered to be the center of the cartilage disc.</li>
		<li>Run a scanner approach and, once the surface of the cartilage disc is reached, retract the cantilever by 100 &#181;m.</li>
	</ol>
	</li>
	<li>Force-distance curve measurements
	<ol>
		<li>Focus on the cells positioned in the desired measurement site. Click the Run button to start the measurements and the generation of force-distance curves in the targeted position.</li>
		<li>Acquire five force-distance curves on each measurement site. Retract the cantilever by 500 &#181;m and move the cantilever to the next measuring site. 
		NOTE: The retraction of the cantilever is a crucial step, as the cartilage disc surface is not homogenous and has irregularities. A high hillock on the surface of the sample can result in a dramatic collision, leading to unwanted cantilever tip/sample damage. We recommend selecting a minimum of five different measurement sites dispersed across the surface of the disc and acquiring a minimum of five force-distance curves at each site.</li>
		<li>Inspect the force-distance curves and save them.</li>
	</ol>
	</li>
	<li>Estimation of Young&#39;s moduli using the Hertz fit model
	<ol>
		<li>Open the generated force-distance curves to be analyzed (.jpk file) in the data analysis software using the Open a Batch of Spectroscopy Curves option.</li>
		<li>Select the Hertz fit model and then select the Elasticity fit option.
		<ol>
			<li>The elasticity fit option automatically performs the following computations on the selected force-distance curve: calculates the baseline and subtracts from the whole curve to remove the baseline offset (the baseline is brought back to zero on the y-axis); determines the contact point by detecting the point where the force-distance curve crosses the zero force-line (the contact point is set to zero on the x-axis); calculates the tip-sample separation (the height signal of the piezo accounting for the bending of the cantilever is subtracted); and fits the force-distance curve automatically with the selected model. If desired, each of these steps can also be carried out independently.</li>
		</ol>
		</li>
		<li>Adapt using the following fit parameters: Poisson ratio of 0.5 and the appropriate cantilever tip radius. 
		NOTE: When using a cantilever with a spherical cantilever tip, the Hertz fit model should be used. The cantilever used in this study had a spherical tip with a radius of 5 &#181;m. We recommend fitting the force-distance curve until the maximum applied force is reached (setpoint).</li>
		<li>Check visually the force-distance curve fit to ensure correctness. This step has to be done for each of the force-distance curves analyzed.</li>
	</ol>
	</li>
	<li>Indentation depth determination 
	NOTE: Depending on the data analysis tool being used, this process may differ. The experimenter can easily read the indentation depth by following a series of steps that are included in the data analysis program.
	<ol>
		<li>Open each of the generated force-distance curves in the data analysis software and select the Hertz fit Model as the analysis process.</li>
		<li>Apply the Subtract Baseline Offset option to zero the vertical deflection axis (y-axis) and select the Offset + Tilt function.</li>
		<li>Use the Find Contact Point function to automatically identify the contact point, which is automatically brought to an x-coordinate of zero.</li>
		<li>Subtract the distance accounting solely for cantilever deflection from the raw piezo height during the indentation using the Vertical Tip Position function.</li>
		<li>Select the Elasticity Fit option to display the processed force-distance curve and select the area of the graph so that it lines up with the most negative value on the Vertical Tip Position Axis (x-axis).</li>
		<li>Read and document the indentation from the X Min box in the parameter tab. Save and document the results.</li>
	</ol>
	</li>
</ol>
4. Statistical analysis
<ol>
	<li>Open the statistical software. Select New Dataset from the drop-down menu.</li>
	<li>Open the Variable View tab after selecting the DataSet file. Define the numerical variables for each cellular pattern category: single strings = SS, double strings = DS, small clusters = SC, big clusters = BC, diffuse and the Young&#39;s moduli.</li>
	<li>In the data view tab, enter the measured Young&#39;s moduli data for each of the corresponding cellular pattern categories. Analyze the data distribution by selecting Analyze from the menu bar, and then Exploratory Data Analysis.</li>
	<li>Select Young&#39;s Moduli as the dependent variable and Cellular Pattern as the factor list. A box plot used for the results section is displayed among the results in the output file.</li>
	<li>To conduct a statistical analysis, choose Dependent Samples in the nonparametric test section of the analyze menu bar tab. Select Young&#39;s Moduli as Test Fields and Cellular Pattern as Groups under the fields tab. Press Run. 
	NOTE: The results are displayed in the output file. For the statistical analysis, a Friedman test is performed.</li>
	<li>Incorporate the nonparametric test&#39;s p-values into the box plot that was created in step 4.4. Save the results by clicking File in the menu bar and selecting Save.</li>
</ol>

The authors have nothing to disclose.

As a progressive and multifactorial disease, OA triggers structural and functional changes in the articular cartilage.Throughout the course of OA, impairments in mechanical features are accompanied by structural and biochemical changes at the surface of the articular cartilage27,31. The earliest pathological events occurring in OA are proteoglycan depletion coupled with collagen network disruption32,33<su...

Due to the ever-shrinking size of electronic devices and systems, the rapid development of micro- and nano-based technology and equipment has gained momentum. One such device is atomic force microscopy (AFM), which can scan biological surfaces and retrieve topographic or biomechanical information at both nano- and micrometer scales1,2. Among its vast features, this tool can be operated as a micro- as well as a nano-indenter to obtain information about the mechanical properties of various biological systems3,4,5,6. The data are collected by physical contact with the surface through a mechanical probe, which can be as small as about 1 nm at its tip7. The resulting deformation of the sample is then displayed based on the indentation depth of the cantilever tip and the force applied on the sample8.
Osteoarthritis (OA) is a long-term degenerative chronic disease characterized by deterioration of the articular cartilage in the joints and surrounding tissues, which can lead to complete exposure of the bone surfaces. The burden of OA is substantial; currently, half of all women and one-third of all men aged 65 and over suffer from OA9. Traumas, obesity, and the resulting altered biomechanics of the joint10&#160;determine the articular cartilage degeneration, which is viewed as a common end result. The pioneering study of Ganz et al. posited that the early steps of the OA process may involve the biomechanical properties of cartilage11, and since then researchers have confirmed this hypothesis12. Likewise, it is generally accepted that the biomechanical properties of the tissue are functionally orchestrated by the ultrastructural organization as well as cell-cell and cell-matrix crosstalk. Any alterations can dramatically impact the overall tissue biomechanical functioning13. To date, OA diagnosis is clinical and is based on plain film radiography14. This approach is two-sided: firstly, the lack of a defined degenerative cut-off threshold to formulate the diagnosis of OA makes the condition difficult to quantify, and, secondly, imaging methods lack sensitivity and standardization and cannot detect localized cartilage damage15,16,17. To this end, the assessment of the mechanical properties of the cartilage has the decisive advantage that it describes a parameter that changes during the course of OA regardless of the etiology of the disease and has a direct influence on tissue functionality at a very early stage. Indentation instruments measure the force by which the tissue resists the indentation. This is, in fact, not a new concept; the earliest studies date back to the 1980s and 1990s. In this period, numerous studies suggested that indentation instruments designed for the arthroscopic measurements of articular cartilage could be well suited to detect degenerative changes in the cartilage. Even 30 years ago, some studies were able to demonstrate that indentation instruments were able to detect in vivo changes in the cartilage surface during tissue degeneration by conducting compressive stiffness measurements during arthroscopy18,19,20.
AFM indentation (AFM-IT) of the articular cartilage provides information about a pivotal mechanical property of the tissue, namely, stiffness. This is a mechanical parameter that describes the relation between an applied, nondestructive load and the resultant deformation of the indented tissue area21. AFM-IT has been shown to be capable of quantifying age-dependent modifications in stiffness in macroscopically unaffected collagen networks, thus, differentiating between the pathological changes associated with OA onset (grade 0 on the Outerbridge scale in articular cartilage)22. We have previously shown that AFM-ITs, on the basis of spatial chondrocyte organization as an image-based biomarker for early cartilage degeneration, allow for not only quantifying but also actually pinpointing the earliest degenerative mechanical changes. These findings have already been confirmed by others23,24. Hence, AFM-IT acts as an interesting tool to diagnose and identify early degenerative changes. These changes can be already measured at a cellular level, reshaping the understanding of the OA pathophysiological process.
In this protocol, we demonstrate a complete histological and biomechanical grading procedure of articular cartilage explants, from native cartilage explant preparation to AFM data acquisition and processing. Through a step-by-step approach, we show how to generate, grade, and visually classify articular cartilage tissue according to different stages of degeneration by means of 2D large mosaic imaging, followed by micro-AFM indentations.
Even though, currently, AFM-IT is one of the most sensitive tools to measure biomechanical changes in cartilage7, like any other instrumental technique, it has limitations and practical peculiarities25&#160;that can lead to erroneous data acquisition. To this end, we subject to scrutiny the most common problems that arise during AFM measurements of the cartilage explants and describe, where possible, how to minimize or overcome them. These include topographical aspects of the samples and the difficulties to stabilize them in an AFM-compatible environment, physical peculiarities of the tissue&#39;s surface, and the resulting difficulties in performing AFM measurements on such surfaces. Examples of erroneous force-distance curves are also presented, emphasizing the conditions that may cause them. Additional limitations inherent to the geometry of the cantilever tip and the use of the Hertz model for the data analysis are also discussed.

<table><tbody><tr><td>Amphotericin B</td><td>Merck KGaA, Darmstadt, Germany</td><td>1397-89-3</td><td></td></tr><tr><td>Atomic force microscop (AFM) head&amp;nbsp;</td><td>CellHesion 200, Bruker Nano GmbH, Berlin, Germany</td><td>JPK00518</td><td></td></tr><tr><td>Biocompatible sample glue&amp;nbsp;</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>H000033</td><td></td></tr><tr><td>Calcein AM</td><td>Cayman, Ann Arbor, Michigan, USA</td><td>14948</td><td>Cell membrane permeable stain, used for cartilage disc sorting- top view imaging</td></tr><tr><td>Cantilever</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>SAA-SPH-5UM</td><td>Frequency Nom: 30KHz, k: 0.2N/m, lenght nom: 115&amp;mu;m, width nom: 40&amp;mu;m,&amp;nbsp; geometry: rectangular, cylindrical tip with a 5&amp;mu;m end radius</td></tr><tr><td>Cartilage ctting device&amp;nbsp;</td><td>Self-made&amp;nbsp;</td><td>n/a</td><td>Cutting plastic device containing predefined wholes of 4mmx1mm</td></tr><tr><td>CDD camera integrated in the AFM</td><td>The Imaging Source Europe GmbH, Bremen, Germany</td><td>DFK 31BF03</td><td></td></tr><tr><td>CDD camera integrated in the fluorescence microscope</td><td>Leica Biosystems, Wetzlar, Germany</td><td>DFC3000G</td><td></td></tr><tr><td>Cryotome</td><td>Leica Biosystems, Wetzlar, Germany</td><td>CM3050S&amp;nbsp;</td><td></td></tr><tr><td>Data Processing Software for the AFM</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>n/a</td><td>Version 5.0.86,&amp;nbsp; can be downloaded for free from the following website https://customers.jpk.com</td></tr><tr><td>Dulbecco&#039;s modified Eagle&#039;s medium (DMEM)&amp;nbsp;</td><td>Gibco, Life Technologies, Darmstadt, Germany</td><td>41966052</td><td></td></tr><tr><td>Fluorescence Microscope (Leica DMi8)</td><td>Leica Biosystems, Wetzlar, Germany</td><td>11889113</td><td></td></tr><tr><td>Glass block cantiliver holder</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>SP-90-05</td><td>Extra long glass block with angled faces, designed especially for the use with the JPK PetriDishHeaterTM (Bruker).</td></tr><tr><td>Inverted phase contrast microscope (integrated in the AFM)</td><td>AxioObserver D1, Carl Zeiss Microscopy, Jena, Germany</td><td>L201306_03</td><td></td></tr><tr><td>Leibovitz&#039;s L-15 medium without L-glutamine&amp;nbsp;</td><td>Merck KGaA, Darmstadt, Germany</td><td>F1315</td><td></td></tr><tr><td>Microscope glass slides</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>CLS294775X50</td><td></td></tr><tr><td>Mounting medium With DAPI</td><td>ibidi GmbH, Gr&amp;auml;felfing, Germany</td><td>50011</td><td>Mounting media with nuclear DAPI (4&amp;prime;,6-diamidino-2-phenylindole) counterstaining used for cartilage discs&amp;nbsp; side view imaging</td></tr><tr><td>Penicillin-Streptomycin</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>P4333</td><td></td></tr><tr><td>Petri dish heater associated with AFM (Petri Dish Heater)</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>T-05-0117</td><td></td></tr><tr><td>Scalpel</td><td>Feather Medical Products, Osaka, Japan</td><td>2023-01</td><td></td></tr><tr><td>Silicone Skirt</td><td>Bruker Nano GmbH, Berlin, Germany</td><td>n/a</td><td>Protective silicone membrane (D55x0.25) which is placed on the basis of the base of the glas block to prevent&amp;nbsp; medium condensation in the AFM head.</td></tr><tr><td>Statistical program - SPSS</td><td>IBM, Armonk, New York, USA</td><td>SPSS Statistics 22</td><td>Vesion 280.0.0.0 (190)</td></tr><tr><td>Tissue culture dishes&amp;nbsp;</td><td>TPP Techno Plastic Products AG, Trasadingen, Switzerland</td><td>TPP93040</td><td></td></tr><tr><td>Tissue-tek O.C.T. Compound</td><td>Sakura Finetek, Alphen aan den Rijn, Netherlands</td><td>SA6255012</td><td>Water-soluble embedding medium&amp;nbsp;</td></tr></tbody></table>

addressing practical issues in atomic force microscopy-based micro-indentation on human articular cartilage explants

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

Using a self-made cutting device, we were able to explant and generate small (4 mm x 1 mm) cartilage discs from fresh human condyles containing a single cellular spatial pattern30&#160;of single strings (SS, Figure 2A), double strings (DS), small clusters (SC), big clusters (BC; Figure 2A), and diffuse (Figure 2B). A representative cartilage explant is depicted in Figure 3A. The ...

Watch this Scientific Journal Video about Addressing Practical Issues in Atomic Force Microscopy-Based Micro-Indentation on Human Articular Cartilage Explants at JoVE.com

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

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

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

addressing-practical-issues-atomic-force-microscopy-based-micro

Research

JoVE Journal

Bioengineering

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

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

Quantitative Hardness Measurement by Instrumented AFM-indentation

In this work, a combination of amplitude-modulated non-contact atomic force microscopy and atomic force spectroscopy is applied for instrumented hardness measurements on an Au(111) surface with atomistic resolution of single plasticity events. A careful experimental procedure is described that includes the force sensor selection, its calibration, the calibration of the cantilever deflection detection system, and the minimization of instrumental drift for accurate and reproducible force-distance measurements. Also, a method for the data analysis is presented that allows the extraction of force-penetration curves from recorded force-distance curves. A typical curve displays a clear elastic deformation regime up to the first plasticity event, or pop-in, with a length in the range of one to two Burger's vectors. Later plasticity events exhibit the same magnitude. The work of plasticity is further extracted from the measurements. Finally, the hardness is determined in combination with the indentation curve using non-contact atomic force microscopy images of the remaining indents.

This experimental protocol describes how atomic force microscopy can be used to measure hardness at the true nanometer scale and to detect single atomistic plasticity events.

In this work, a combination of amplitude-modulated non-contact atomic force microscopy and atomic force spectroscopy is applied for instrumented hardness ...

Engineering

Biomechanical Characterization of Human Soft Tissues Using Indentation and Tensile Testing

Regenerative medicine aims to engineer materials to replace or restore damaged or diseased organs. The mechanical properties of such materials should mimic the human tissues they are aiming to replace; to provide the required anatomical shape, the materials must be able to sustain the mechanical forces they will experience when implanted at the defect site. Although the mechanical properties of tissue-engineered scaffolds are of great importance, many human tissues that undergo restoration with engineered materials have not been fully biomechanically characterized. Several compressive and tensile protocols are reported for evaluating materials, but with large variability it is difficult to compare results between studies. Further complicating the studies is the often destructive nature of mechanical testing. Whilst an understanding of tissue failure is important, it is also important to have knowledge of the elastic and viscoelastic properties under more physiological loading conditions.
This report aims to provide a minimally destructive protocol to evaluate the compressive and tensile properties of human soft tissues. As examples of this technique, the tensile testing of skin and the compressive testing of cartilage are described. These protocols can also be directly applied to synthetic materials to ensure that the mechanical properties are similar to the native tissue. Protocols to assess the mechanical properties of human native tissue will allow a benchmark by which to create suitable tissue-engineered substitutes.

Tissue biomechanics is important for maintaining cell shape and function and for determining phenotype. This report demonstrates non-destructive mechanical protocols for characterizing elastic and viscoelastic properties of human soft tissues, which can be directly applied to tissue-engineered substrates to allow a close matching of engineered materials to native tissue.

Regenerative medicine aims to engineer materials to replace or restore damaged or diseased organs. The mechanical properties of such materials should ...

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 ...

Real-time Visualization and Analysis of Chondrocyte Injury Due to Mechanical Loading in Fully Intact Murine Cartilage Explants

Homeostasis of articular cartilage depends on the viability of resident cells (chondrocytes). Unfortunately, mechanical trauma can induce widespread chondrocyte death, potentially leading to irreversible breakdown of the joint and the onset of osteoarthritis. Additionally, maintenance of chondrocyte viability is important in osteochondral graft procedures for optimal surgical outcomes. We present a method to assess the spatial extent of cell injury/death on the articular surface of intact murine synovial joints after application of controlled mechanical loads or impacts. This method can be used in comparative studies to investigate the effects of different mechanical loading regimens, different environmental conditions or genetic manipulations, as well as different stages of cartilage degeneration on short- and/or long-term vulnerability of in situ articular chondrocytes. The goal of the protocol introduced in the manuscript is to assess the spatial extent of cell injury/death on the articular surface of murine synovial joints. Importantly, this method enables testing on fully intact cartilage without compromising native boundary conditions. Moreover, it allows for real-time visualization of vitally stained articular chondrocytes and single image-based analysis of cell injury induced by application of controlled static and impact loading regimens. Our representative results demonstrate that in healthy cartilage explants, the spatial extent of cell injury depends sensitively on load magnitude and impact intensity. Our method can be easily adapted to investigate the effects of different mechanical loading regimens, different environmental conditions or different genetic manipulations on the mechanical vulnerability of in situ articular chondrocytes.

We present a method to assess the spatial extent of cell injury/death on the articular surface of intact murine joints after application of controlled mechanical loads or impacts. This method can be used to investigate how osteoarthritis, genetic factors and/or different loading regimens affect the vulnerability of in situ chondrocytes.

Homeostasis of articular cartilage depends on the viability of resident cells (chondrocytes). Unfortunately, mechanical trauma can induce widespread ...

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 ...

Biology

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 ...

Injection of Porcine Adipose Tissue-Derived Stroma Cells via Waterjet Technology

Urinary incontinence (UI) is a highly prevalent condition characterized by the deficiency of the urethral sphincter muscle. Regenerative medicine branches, particularly cell therapy, are novel approaches to improve and restore the urethral sphincter function. Even though injection of active functional cells is routinely performed in clinical settings by needle and syringe, these approaches have significant disadvantages and limitations. In this context, needle-free waterjet (WJ) technology is a feasible and innovative method that can inject viable cells by visual guided cystoscopy in the urethral sphincter. In the present study, we used WJ to deliver porcine adipose tissue-derived stromal cells (pADSCs) into cadaveric urethral tissue and subsequently investigated the effect of WJ delivery on cell yield and viability. We also assessed the biomechanical features (i.e., elasticity) by atomic force microscopy (AFM) measurements. We showed that WJ delivered pADSCs were significantly reduced in their cellular elasticity. The viability was significantly lower compared to controls but is still above 80%.

We present a method of cell injection via needle free waterjet technology coupled with a sequela of post-delivery investigations in terms of cellular viability, proliferation, and elasticity measurements.

Urinary incontinence (UI) is a highly prevalent condition characterized by the deficiency of the urethral sphincter muscle. Regenerative medicine ...

Experimental and Data Analysis Workflow for Soft Matter Nanoindentation

Nanoindentation refers to a class of experimental techniques where a micrometric force probe is used to quantify the local mechanical properties of soft biomaterials and cells. This approach has gained a central role in the fields of mechanobiology, biomaterials design&#160;and tissue engineering, to obtain a proper mechanical characterization of soft materials with a resolution comparable to the size of single cells (&#956;m). The most popular strategy to acquire such experimental data is to employ an atomic force microscope (AFM); while this instrument offers an unprecedented resolution in force (down to pN) and space (sub-nm), its usability is often limited by its complexity that prevents routine measurements of integral indicators of mechanical properties, such as Young&#39;s Modulus (E). A new generation of nanoindenters, such as those based on optical fiber sensing technology, has recently gained popularity for its ease of integration while allowing to apply sub-nN forces with &#181;m spatial resolution, therefore being suitable to probe local mechanical properties of hydrogels and cells.
In this protocol, a step-by-step guide detailing the experimental procedure to acquire nanoindentation data on hydrogels and cells using a commercially available ferrule-top optical fiber sensing nanoindenter is presented. Whereas some steps are specific to the instrument used herein, the proposed protocol can be taken as a guide for other nanoindentation devices, granted some steps are adapted according to the manufacturer&#39;s guidelines. Further, a new open-source Python software equipped with a user-friendly graphical user interface for the analysis of nanoindentation data is presented, which allows for screening of incorrectly acquired curves, data filtering, computation of the contact point through different numerical procedures, the conventional computation of E, as well as a more advanced analysis particularly suited for single-cell nanoindentation data.

The protocol presents a complete workflow for soft material nanoindentation experiments, including hydrogels and cells. First, the experimental steps to acquire force spectroscopy data are detailed; then, the analysis of such data is detailed through a newly developed open-source Python software, which is free to download from GitHub.

Nanoindentation refers to a class of experimental techniques where a micrometric force probe is used to quantify the local mechanical properties of soft ...

A Probing Device for Quantitatively Measuring the Mechanical Properties of Soft Tissues during Arthroscopy

Probing in arthroscopic surgery is performed by pulling or pushing the soft tissue, which provides feedback for understanding the condition of the soft tissue. However, the output is only qualitative based on the &#8220;surgeon&#8217;s feeling&#8221;. Herein is described a probing device developed to address this issue by measuring the resistance of soft tissues quantitatively with a tri-axial force sensor. Under both conditions (i.e., pull- and push-probing certain tissues mimicking the acetabular labrum and cartilage), this probing device is found to be useful for measuring some mechanical properties in joints during arthroscopy.

Probing during arthroscopy surgery is normally done to assess the condition of the soft tissue, but this approach has always been subjective and qualitative. This report describes a probing device that can measure the resistance of the soft tissue quantitatively with a tri-axial force sensor during arthroscopy.

Probing in arthroscopic surgery is performed by pulling or pushing the soft tissue, which provides feedback for understanding the condition of the soft ...

Medicine

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

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

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