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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;</td>
			<td>CellHesion 200, Bruker Nano GmbH, Berlin, Germany</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>Biocompatible sample glue&#160;</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&#956;m, width nom: 40&#956;m,&#160; geometry: rectangular, cylindrical tip with a 5&#956;m end radius</td>
		</tr>
		<tr>
			<td>Cartilage ctting device&#160;</td>
			<td>Self-made&#160;</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&#160;</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,&#160; can be downloaded for free from the following website https://customers.jpk.com</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified Eagle&#39;s medium (DMEM)&#160;</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&#39;s L-15 medium without L-glutamine&#160;</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&#228;felfing, Germany</td>
			<td>50011</td>
			<td>Mounting media with nuclear DAPI (4&#8242;,6-diamidino-2-phenylindole) counterstaining used for cartilage discs&#160; 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&#160; 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&#160;</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&#160;</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

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

891 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

Bacterial Immobilization for Imaging by Atomic Force Microscopy

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface. It has the ability to image cells and biomolecules, in a liquid environment, without the need to chemically treat the sample. In order to accomplish this goal, the sample must sufficiently adhere to the mounting surface to prevent removal by forces exerted by the scanning AFM cantilever tip. In many instances, successful imaging depends on immobilization of the sample to the mounting surface. Optimally, immobilization should be minimally invasive to the sample such that metabolic processes and functional attributes are not compromised. By coating freshly cleaved mica surfaces with porcine (pig) gelatin, negatively charged bacteria can be immobilized on the surface and imaged in liquid by AFM. Immobilization of bacterial cells on gelatin-coated mica is most likely due to electrostatic interaction between the negatively charged bacteria and the positively charged gelatin. Several factors can interfere with bacterial immobilization, including chemical constituents of the liquid in which the bacteria are suspended, the incubation time of the bacteria on the gelatin coated mica, surface characteristics of the bacterial strain and the medium in which the bacteria are imaged. Overall, the use of gelatin-coated mica is found to be generally applicable for imaging microbial cells.

Live Gram-negative and Gram-positive bacteria can be immobilized on gelatin-coated mica and imaged in liquid using Atomic Force Microscopy (AFM).

AFM is a high-resolution (nm scale) imaging tool that mechanically probes a surface.  It has the ability to image cells and biomolecules, in a liquid ...

Visualization of Recombinant DNA and Protein Complexes Using Atomic Force Microscopy

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On the end of the microscope's cantilever is a nano-scale probe, which traverses image areas ranging from nanometers to micrometers, measuring the elevation of macromolecules resting on the substrate surface at any given point. Electrostatic forces cause proteins, lipids, and nucleic acids to loosely attach to the substrate in random orientations and permit imaging. The generated data resemble a topographical map, where the macromolecules resolve as three-dimensional particles of discrete sizes (Figure 1) 1,2. Tapping mode AFM involves the repeated oscillation of the cantilever, which permits imaging of relatively soft biomaterials such as DNA and proteins. One of the notable benefits of AFM over other nanoscale microscopy techniques is its relative adaptability to visualize individual proteins and macromolecular complexes in aqueous buffers, including near-physiologic buffered conditions, in real-time, and without staining or coating the sample to be imaged. 
The method presented here describes the imaging of DNA and an immunoadsorbed transcription factor (i.e. the glucocorticoid receptor, GR) in buffered solution (Figure 2). Immunoadsorbed proteins and protein complexes can be separated from the immunoadsorbing antibody-bead pellet by competition with the antibody epitope and then imaged (Figure 2A). This allows for biochemical manipulation of the biomolecules of interest prior to imaging. Once purified, DNA and proteins can be mixed and the resultant interacting complex can be imaged as well. Binding of DNA to mica requires a divalent cation 3,such as Ni2+ or Mg2+, which can be added to sample buffers yet maintain protein activity. Using a similar approach, AFM has been utilized to visualize individual enzymes, including RNA polymerase 4 and a repair enzyme 5, bound to individual DNA strands. These experiments provide significant insight into the protein-protein and DNA-protein biophysical interactions taking place at the molecular level. Imaging individual macromolecular particles with AFM can be useful for determining particle homogeneity and for identifying the physical arrangement of constituent components of the imaged particles. While the present method was developed for visualization of GR-chaperone protein complexes 1,2 and DNA strands to which the GR can bind, it can be applied broadly to imaging DNA and protein samples from a variety of sources.

A tapping mode atomic force microscope (AFM) method for the visualization of plasmid DNA, cytoplasmic proteins, and DNA-protein complexes is described. The method includes alternate approaches for preparing samples for AFM imaging following biochemical manipulation. DNA containing specific protein interacting regions are observed in near-physiologic buffer conditions.

Atomic force microscopy (AFM) allows for the visualizing of individual proteins, DNA molecules, protein-protein complexes, and DNA-protein complexes. On ...

Quantifying the Mechanical Properties of the Endothelial Glycocalyx with Atomic Force Microscopy

Our understanding of the interaction of leukocytes and the vessel wall during leukocyte capture is limited by an incomplete understanding of the mechanical properties of the endothelial surface layer. It is known that adhesion molecules on leukocytes are distributed non-uniformly relative to surface topography 3, that topography limits adhesive bond formation with other surfaces 9, and that physiological contact forces (&asymp; 5.0 &minus; 10.0 pN per microvillus) can compress the microvilli to as little as a third of their resting length, increasing the accessibility of molecules to the opposing surface 3, 7. We consider the endothelium as a two-layered structure, the relatively rigid cell body, plus the glycocalyx, a soft protective sugar coating on the luminal surface 6. It has been shown that the glycocalyx can act as a barrier to reduce adhesion of leukocytes to the endothelial surface 4. In this report we begin to address the deformability of endothelial surfaces to understand how the endothelial mechanical stiffness might affect bond formation. Endothelial cells grown in static culture do not express a robust glycocalyx, but cells grown under physiological flow conditions begin to approximate the glycocalyx observed in vivo 2. The modulus of the endothelial cell body has been measured using atomic force microscopy (AFM) to be approximately 5 to 20 kPa 5. The thickness and structure of the glycocalyx have been studied using electron microscopy 8, and the modulus of the glycocalyx has been approximated using indirect methods, but to our knowledge, there have been no published reports of a direct measurement of the glycocalyx modulus in living cells. In this study, we present indentation experiments made with a novel AFM probe on cells that have been cultured in conditions to maximize their glycocalyx expression to make direct measurements of the modulus and thickness of the endothelial glycocalyx.

The mechanical characteristics of endothelial glycocalyx were measured by indentation using micron sized spheres on AFM cantilevers. Endothelial cells were cultured in a custom chamber under physiological flow conditions to induce glycocalyx expression. Data were analyzed using a thin film model to determine the glycocalyx thickness and modulus.

Our  understanding of the interaction of leukocytes and the vessel wall during  leukocyte capture is limited by an incomplete understanding of the ...

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, ...

Human Cartilage Tissue Fabrication Using Three-dimensional Inkjet Printing Technology

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and regenerative medicine. With digital control&#160;cells, scaffolds, and growth factors can be precisely deposited to the desired two-dimensional (2D) and three-dimensional (3D) locations rapidly. Therefore, this technology is an ideal approach to fabricate tissues mimicking their native anatomic structures. In order to engineer cartilage with native zonal organization, extracellular matrix composition (ECM), and mechanical properties, we developed a bioprinting platform using a commercial inkjet printer with simultaneous photopolymerization capable for 3D cartilage tissue engineering. Human chondrocytes suspended in poly(ethylene glycol) diacrylate (PEGDA) were printed for 3D neocartilage construction via layer-by-layer assembly. The printed cells were fixed at their original deposited positions, supported by the surrounding scaffold in simultaneous photopolymerization. The mechanical properties of the printed tissue were similar to the native cartilage. Compared to conventional tissue fabrication, which requires longer UV exposure, the viability of the printed cells with simultaneous photopolymerization was significantly higher. Printed neocartilage demonstrated excellent glycosaminoglycan (GAG) and collagen type II production, which was consistent with gene expression. Therefore, this platform is ideal for accurate cell distribution and arrangement for anatomic tissue engineering.

The methods described in this paper show how to convert a commercial inkjet printer into a bioprinter with simultaneous UV polymerization. The printer is capable of constructing 3D tissue structure with cells and biomaterials. The study demonstrated here constructed a 3D neocartilage.

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and ...

Novel Atomic Force Microscopy Based Biopanning for Isolation of Morphology Specific Reagents against TDP-43 Variants in Amyotrophic Lateral Sclerosis

Because protein variants play critical roles in many diseases including TDP-43 in Amyotrophic Lateral Sclerosis (ALS), alpha-synuclein in Parkinson&#8217;s disease and beta-amyloid and tau in Alzheimer&#8217;s disease, it is critically important to develop morphology specific reagents that can selectively target these disease-specific protein variants to study the role of these variants in disease pathology and for potential diagnostic and therapeutic applications. We have developed novel atomic force microscopy (AFM) based biopanning techniques that enable isolation of reagents that selectively recognize disease-specific protein variants. There are two key phases involved in the process, the negative and positive panning phases. During the negative panning phase, phages that are reactive to off-target antigens are eliminated through multiple rounds of subtractive panning utilizing a series of carefully selected off-target antigens. A key feature in the negative panning phase is utilizing AFM imaging to monitor the process and confirm that all undesired phage particles are removed. For the positive panning phase, the target antigen of interest is fixed on a mica surface and bound phages are eluted and screened to identify phages that selectively bind the target antigen. The target protein variant does not need to be purified providing the appropriate negative panning controls have been used. Even target protein variants that are only present at very low concentrations in complex biological material can be utilized in the positive panning step. Through application of this technology, we acquired antibodies to protein variants of TDP-43 that are selectively found in human ALS brain tissue. We expect that this protocol should be applicable to generating reagents that selectively bind protein variants present in a wide variety of different biological processes and diseases.

Using atomic force microscopy in combination with biopanning technology we created a negative and positive biopanning system to acquire antibodies against disease-specific protein variants present in any biological material, even at low concentrations. We were successful in obtaining antibodies to TDP-43 protein variants involved in Amyotrophic Lateral Sclerosis.

Because protein variants play critical roles in many diseases including TDP-43 in Amyotrophic Lateral Sclerosis (ALS), alpha-synuclein in ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

3D Hydrogel Scaffolds for Articular Chondrocyte Culture and Cartilage Generation

Human articular cartilage is highly susceptible to damage and has limited self-repair and regeneration potential. Cell-based strategies to engineer cartilage tissue offer a promising solution to repair articular cartilage. To select the optimal cell source for tissue repair, it is important to develop an appropriate culture platform to systematically examine the biological and biomechanical differences in the tissue-engineered cartilage by different cell sources. Here we applied a three-dimensional (3D) biomimetic hydrogel culture platform to systematically examine cartilage regeneration potential of juvenile, adult, and osteoarthritic (OA) chondrocytes. The 3D biomimetic hydrogel consisted of synthetic component poly(ethylene glycol) and bioactive component chondroitin sulfate, which provides a physiologically relevant microenvironment for in vitro culture of chondrocytes. In addition, the scaffold may be potentially used for cell delivery for cartilage repair in vivo. Cartilage tissue engineered in the scaffold can be evaluated using quantitative gene expression, immunofluorescence staining, biochemical assays, and mechanical testing. Utilizing these outcomes, we were able to characterize the differential regenerative potential of chondrocytes of varying age, both at the gene expression level and in the biochemical and biomechanical properties of the engineered cartilage tissue. The 3D culture model could be applied to investigate the molecular and functional differences among chondrocytes and progenitor cells from different stages of normal or aberrant development.

Cartilage repair represents an unmet medical challenge and cell-based approaches to engineer human articular cartilage are a promising solution. Here, we describe three-dimensional (3D) biomimetic hydrogels as an ideal tool for the expansion and maturation of human articular chondrocytes.

Human articular cartilage is highly susceptible to damage and has limited self-repair and regeneration potential. Cell-based strategies to engineer ...

An Experimental and Finite Element Protocol to Investigate the Transport of Neutral and Charged Solutes across Articular Cartilage

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular cartilage impairs its load-bearing function substantially as it experiences tremendous chemical degradation, i.e. proteoglycan loss and collagen fibril disruption. One promising way to investigate chemical damage mechanisms during OA is to expose the cartilage specimens to an external solute and monitor the diffusion of the molecules. The degree of cartilage damage (i.e. concentration and configuration of essential macromolecules) is associated with collisional energy loss of external solutes while moving across articular cartilage creates different diffusion characteristics compared to healthy cartilage. In this study, we introduce a protocol, which consists of several steps and is based on previously developed experimental micro-Computed Tomography (micro-CT) and finite element modeling. The transport of charged and uncharged iodinated molecules is first recorded using micro-CT, which is followed by applying biphasic-solute and multiphasic finite element models to obtain diffusion coefficients and fixed charge densities across cartilage zones.

We propose a protocol to investigate the transport of charged and uncharged molecules across articular cartilage with the aid of recently developed experimental and numerical methods.

Osteoarthritis (OA) is a debilitating disease that is associated with degeneration of articular cartilage and subchondral bone. Degeneration of articular ...

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 ...