We thank our co-authors from the original publication for their help and support.

The human cartilage samples were obtained from patients undergoing total knee arthroplasty in the Department of Orthopaedic Surgery of the University Hospital of Tuebingen, Germany, and the Winghofer-hospital, Rottenburg a.N., Germany, for end-stage OA of the knee. Full departmental, institutional, and local ethical committee approval were obtained before commencement of the study (project number 674/2016BO2). Written informed consent was received from all patients before participation. The methods were carried out in accordance with the approved guidelines.
1. Sample preparation
<ol>
	<li>Preparing the cartilage for cryotome sectioning
	<ol>
		<li>To evaluate degenerative changes in the knee joint, take articular cartilage samples obtained from the load-bearing zone of the femoral condyles after tissue resection from the patients. 
		NOTE: Samples that are investigated should still contain at least a thin layer of subchondral bone to allow clear identification of the tissue orientation with respect to the inner and outer surface, thus also allowing a standardized tissue harvest of the topmost cartilage layer. The cartilage can be used for AFM measurements until 24 h after surgery. The tissue can be stored in serum-free Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) with 2% (v/v) penicillin-streptomycin and 1.2% (v/v) amphotericin B prior to use. Storing the samples for more than 24 h can cause artifacts in the AFM measurements due to tissue swelling, however.</li>
		<li>Cut the articular cartilage as a whole from the subchondral bone with the help of a scalpel and afterwards embed the cartilage sample in water-soluble embedding medium on the cryotome knob that freezes under low temperature in the cryotome device. 
		NOTE: The frozen medium stabilizes the tissue it envelops. It also results in the freezing of the cartilage sample along with the embedding medium. When cutting the cartilage from the bone, track the orientation of the tissue. The tissue embedded for the cryosections must be placed in such a way that the top layer (i.e., articular surface) of the cartilage faces the blade. Note that the tissue has to be fully covered by the embedding medium.</li>
	</ol>
	</li>
	<li>Cryotome sectioning of the cartilage
	<ol>
		<li>Using a standard cryotome, section the tissue at a thickness of 35 &#181;m from the topmost layer (i.e., articular surface) of the articular cartilage that is embedded in the frozen water-soluble embedding medium. 
		NOTE: In total, up to 300 &#181;m (which corresponds to about nine sections) can be sectioned and used for the analyses. The proposed limit of 300 &#181;m corresponds to the visual penetration depth that can usually be obtained with fluorescence microscopy in hyaline cartilage such as articular cartilage. Thus, it is possible to sort cartilage according to the predominant local spatial pattern and then measure these patterns in the corresponding sections.</li>
		<li>Collect the sections on a glass slide and rinse the sections 3x with phosphate-buffered saline (PBS) to remove the water-soluble embedding medium.</li>
	</ol>
	</li>
	<li>Gluing of cartilage sections onto an AFM compatible Petri dish 
	NOTE: Because the AFM gathers data by mechanical indentation on the sample, the sections need to be fixed in place to allow for precise measurements.
	<ol>
		<li>Gently glue the 35 &#181;m thick sections with biocompatible sample glue onto tissue culture dishes that are compatible with the AFM device. To do this, take 2&#8722;3 drops of the glue and put them on the tissue culture dish at the location where the edge of the section will end up. Now put the cartilage section on the glue and allow it to stick firmly to the surface of the tissue culture dish. 
		NOTE: The 35 &#181;m thick sections of hyaline articular cartilage are not very prone to curling. However, if curling occurs when removing the samples from the buffer medium, make sure that as little fluid as possible sticks to the samples. As soon as the excess water has dried up, directly spread the tissue on the dispersed spots of glue so that it is fixed to the surface of the cell culture dish. The glue is applied in a thin layer only at the edges of the sample and not on the entire section. Measure only that portion of the sample that is far from the glued area to eliminate the possibility of the glue interfering with the measurements.</li>
		<li>Incubate the sections at room temperature (RT) for 2 min to allow the glue and tissue to bond properly. Then cover the sections with Leibovitz&#39;s L-15 medium without L-glutamine and without Sodium bicarbonate so that they are fully submerged. 
		NOTE: Sudden movements of the sample during fixation or insufficient glue application are common mistakes that result in the tissue&#8217;s detachment from the culture dish. Furthermore, applying the Leibowitz medium should be done in a gentle way, so as not to detach the sample from the surface due to &#34;waves&#34; created by the medium.</li>
		<li>Place the Petri dish with the glued sections in the standard cell culture incubator at 37 &#176;C until measurements are performed. 
		NOTE: Make sure that the tissue sections are stable and do not detach from the surface of the tissue culture dish by performing each step slowly.</li>
	</ol>
	</li>
</ol>
2. Cantilever preparation (gluing the microspheres)
<ol>
	<li>Preparing the AFM device and diluting the microspheres
	<ol>
		<li>Place the cantilever on the glass block specified for air measurements, fix it with the spring, and mount the glass block on the AFM head.</li>
		<li>Dilute the microspheres in 100% ethanol (100 particles/10 &#181;L) and ultrasonicate for 10 s, so that the microspheres are separated and do not clump together.</li>
	</ol>
	</li>
	<li>Preparing the setup for gluing
	<ol>
		<li>Clean a glass slide with 70% ethanol and mount it on the sample holder on the AFM device. 
		NOTE: The cleansing of the slide prevents specks of dust that might interfere with the gluing process from accumulating on the slide&#39;s surface.</li>
		<li>Place 2 &#181;L of the microsphere suspension in the middle of the slide and 2 &#181;L of glue near the microsphere suspension. 
		NOTE: Placing the microsphere suspension and glue close to each other is recommended to allow for a quick transition of the cantilever between the glue and the microspheres. If the transition between dipping the cantilever into the glue and making contact with a microsphere is too long, the glue will eventually harden, thereby losing its adhesive properties.</li>
		<li>Let the ethanol liquid in the microsphere suspension air-dry, leaving the microspheres in place.</li>
	</ol>
	</li>
	<li>Gluing the microspheres on the cantilever tip
	<ol>
		<li>Dip the cantilever manually into the glue with the help of the stepper motor in 10 &#181;m steps until the tip is covered by glue. Perform this step swiftly, as the glue dries fast.</li>
		<li>Retract the cantilever again by 100 &#181;m, move it towards the microspheres, and position the cantilever tip directly above a single microsphere.</li>
		<li>Run a force spectroscopy measurement to dip the cantilever tip onto the selected microsphere with the parameters shown in Table 1. 
		NOTE: By executing this step, a measurement of the microsphere&#39;s surface is used to establish contact between the cantilever&#39;s tip and the microsphere. The relatively long contact time shown in Table 1 allows for ample bonding time between the glue and microsphere. The force-distance curve obtained during this step is generated as a by-product of gluing the microsphere on the cantilever tip and is not processed for further analysis.</li>
		<li>Once a microsphere is attached to the cantilever, retract the glass block with the cantilever mounted on top and then remove the AFM head from the microscope. Lastly, unmount the glass block and cantilever from the AFM head.</li>
		<li>Incubate the cantilever at 65 &#176;C for 2 h and let it dry overnight at RT before starting the measurements. 
		NOTE: Incubation of the cantilever enhances the glue&#39;s adhesive properties, rendering the cantilever-microsphere-complex more stable.</li>
	</ol>
	</li>
</ol>
3. Preparing the AFM device for measurements
<ol>
	<li>Adjust a glass block specified for measuring in a liquid environment on the AFM holder so that the upper surface is straight and parallel to the AFM holder.</li>
	<li>With the help of tweezers, carefully mount the selected cantilever on the surface of the glass block, so that the AFM tip with the microsphere protrudes over the polished optical plane. 
	NOTE: The cantilever&#39;s tip needs to protrude over the polished plane in order to be able to reflect the AFM&#39;s laser onto the photodetector. Be very careful when placing the cantilever on the AFM so as not to scratch the polished optical surface of the glass block. Scratching the glass block may result in difficulties in adjusting the laser alignment and may interfere with the subsequent measurements.</li>
	<li>Because the cantilever will be subject to mechanical pressure, it needs to be fixed in place. Stabilize the cantilever on the glass block by sliding the metallic spring into the groove of the block and clamping the top of the cantilever with the spring with the help of tweezers.</li>
	<li>Carefully place the glass block with the cantilever on the AFM head and secure it with the integrated locking mechanism. Make sure that the spring is facing to the left side so that the cantilever is placed in the correct orientation. Then mount the AFM head with the cantilever on the AFM device.</li>
</ol>
4. Loading the sample and calibration of the cantilever
NOTE: Here, calibration of the device is performed by running a force curve on the clean surface of the Petri dish filled with Leibovitz&#8217;s medium without any sample tissue. Calibration can also be performed by using a separate control AFM dish filled only with the AFM medium without the sample.
<ol>
	<li>Mounting the sample on the AFM device
	<ol>
		<li>Place the Petri dish prepared in step 1.3.3 onto the AFM sample holder.</li>
		<li>Turn on the Petri dish heater and set it at 37 &#176;C. Allow the tissue culture dish to reach optimal temperature (i.e., 20 min).</li>
		<li>Place a protecting foil on the base of the cantilever assembly to avoid the condensation of medium in the AFM head.</li>
	</ol>
	</li>
	<li>Calibration of the device
	<ol>
		<li>Open the software (Table of Materials), followed by the laser alignment window, and the window depicting the approach parameter settings. Then turn on the stepper motor, the laser light, and the CCD-camera.</li>
		<li>Use the CCD-camera on the microscope to visualize and identify the cantilever. Using the stepper motor function, lower the cantilever in 100 &#181;m steps until the cantilever is fully submerged in the medium. 
		NOTE: The submerging of the cantilever in the liquid is visually identifiable via the CCD-camera. When it breaks the water&#39;s surface, the cantilever tip creates easily noticeable circular reflections in the water.</li>
		<li>Direct the laser on top of the cantilever using the adjustment screws. 
		NOTE: Do not overwind the screws, as this will damage the laser alignment mechanism. The cantilever is viewed from below and the laser beam comes from above.</li>
		<li>Once the laser is positioned on the cantilever, adjust the laser beam with the help of screws in the AFM device so that the reflected beam falls onto the center of the photodetector. Keep monitoring the position of the laser beam using the laser alignment function. 
		NOTE: The detector picks up the deflection of the laser beam reflected by the cantilever and converts it into an electrical signal.</li>
		<li>After laser-cantilever adjustment, the Sum signal has to be 1 V or above while the lateral and vertical deflections have to be close to 0. If these values are not obtained, adjust the photodetector.</li>
	</ol>
	</li>
	<li>Obtaining the calibration force curve
	<ol>
		<li>In order to reach the surface on the AFM dish to start the measurements, run a scanner Approach with the approach parameters given in Table 2. Once the bottom of the tissue culture dish is reached, retract the cantilever by 100 &#181;m. 
		NOTE: At this point the probe is exactly 100 &#181;m above from the bottom of the tissue culture dish.</li>
		<li>Set up the RUN parameters and adjust the parameters displayed in Table 3. Click the RUN button to start a measurement and obtain a calibration force-distance curve. 
		NOTE: The force curve obtained here is shown in Figure 1.</li>
		<li>On the calibration force-distance curve, select the region for a linear fit of the retracted curve in the software. 
		NOTE: The linear fit is done to measure the cantilever&#8217;s sensitivity. Once the linear fit is in place, the values will be saved by the software. The spring constant measurement or the thermal noise of the cantilever is calculated by the software afterwards.</li>
		<li>As the measurements are performed in medium at a temperature of 37 &#176;C, set the temperature variable at 37 &#176;C in the software to mimic physiological conditions as closely as possible. 
		NOTE: By the end of the calibration, the vertical deflection is saved and displayed in the unit of force, the Newton (N), instead of volts (V), which is the unit of the original registration by the photodiode detector.</li>
	</ol>
	</li>
</ol>
5. Biomechanical characterization of the ECM and PCM by performing elasticity measurements via AFM
<ol>
	<li>Identifying the chondrocyte patterns in the cartilage section
	<ol>
		<li>Observe the cartilage section under a phase contrast microscope integrated in the AFM system and identify the specific cellular patterns10 of articular cartilage from the osteoarthritic knee: single strings (healthy tissue areas), double strings (beginning tissue degeneration), small and big clusters (both advanced tissue degeneration). See Figure 2A&#8211;D.</li>
		<li>Once the specific desired pattern is identified, measure two sites per chosen pattern per matrix type (PCM or ECM) and perform nine measurement repetitions on each measurement site (Figure 2E,F). Ensure a sample size large enough to account for possible inaccuracies. 
		NOTE: For the PCM measurements, place the cantilever in close proximity to the cells (as shown by the red circles in Figure 2E,F). In order to conduct measurements of the ECM, select a region without any cells (shown by the blue area in Figure 2E,F) and perform the indentation as described in step 5.2.</li>
	</ol>
	</li>
	<li>Indentation of the targeted site
	<ol>
		<li>Perform an Approach followed by a retraction so that the cantilever is positioned 100 &#181;m above the tissue, which will be the starting position for the subsequent measurements.</li>
		<li>Focus on the PCM or ECM of the pattern to be measured and fix the computer mouse at that point. 
		NOTE: The mouse will serve as a visual marker for the target site of indentation. Once the focus shifts to the cantilever tip, the tissue structures will become indistinguishable or blurred.</li>
		<li>Now focus on the probe and move the tip of the probe to the point previously fixed by the computer arrow. Next, start the measurements with RUN, using the set point parameter obtained by calibration of the cantilever. 
		NOTE: An exemplary force curve obtained from measuring the PCM of a single string is shown in Supplemental Figure 1.</li>
	</ol>
	</li>
</ol>
6. Data processing
NOTE: The data analysis or determination of the elastic modulus is performed using a Hertz model as described previously11,12. The indenter&#8217;s shape was spherical due to the usage of microspheres on the tip and the Poisson&#8217;s ratio was kept at 0.5 based on previous literature13,14,15.
<ol>
	<li>Open the data processing software (Table of Materials) compatible with the data obtained from the AFM device.</li>
	<li>Select the Hertz model in the software for processing the force-distance curves. Set the Poisson ratio at 0.5 and select the tip shape as spherical and a tip radius of 12.5 &#181;m.</li>
	<li>Once all the parameters are adjusted, the results are fitted, and the Young&#8217;s modulus is calculated and displayed by the software.</li>
</ol>

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The authors have nothing to disclose.

Using AFM as a novel and powerful technique to measure the biomechanical properties of biological materials at a nanoscale level, we measured the elastic properties of the ECM and PCM in human osteoarthritic articular cartilage. Cartilage samples were selected according to their predominant spatial pattern of chondrocyte organization as an image-based biomarker for local tissue degeneration. As expected, a strong decline in the values of elasticity of both ECM and PCM was observed along spatial chondrocyte reorganization. These observations clearly highlight that the deviations in spatial arrangement of chondrocytes were not only associated with changes in the elastic properties of the cellular microenvironment (PCM), but also throughout the entire cartilage (ECM). Furthermore, the ECM/PCM ratio did not show any significant changes in spite of the robust changes in the elastic moduli of PCM and ECM during OA. These findings indicate that the changes in the mechanical properties of the ECM and the PCM occurred unidirectionally and at the same time, which might mean that the nature of progressive destruction is similar for both the PCM and ECM. OA initiation and progression thus triggers significant PCM and ECM degradation and ultimately destruction. Both losses were associated with a significant loss of biomechanical properties of the articular cartilage such as its elasticity, as shown in the present study. This emphasizes the functional relevance of the spatial organization of chondrocytes as a marker for biomechanical properties. Conversely, it allows us to use local elasticity measurements to draw conclusions about the locally predominant spatial patterns and thus the local tissue degeneration of the cartilage.
Atomic force microscopy (AFM) has emerged as a high-resolution tool to study tissues in a nondestructive way. It operates by physically probing samples with a delicate and pliable cantilever that reflects a laser onto a photodiode. Any changes in this reflection are registered and converted into an electrical signal. While AFM is a powerful tool to conduct nanoscale measurements, it does not come without its limitations and pitfalls. Especially critical is the cantilever preparation by gluing the microspheres. In the context of this method, microspheres are used to modify the indentation depth and local pressure during measurements. Using small microspheres attached to the tip of the probe allows the measurement of the biomechanical properties of a fiber network rather than the elasticity of single fiber when using the tip alone. It also prevents tissue damage during the measurement process. Due to the delicate nature of the cantilever sensors, an attentive and careful mode of preparation needs to be established in order to obtain consistent and accurate measurements. In order to prevent the microspheres from detaching from the sensor&#8217;s tip, we recommend freshly mixed glue not older than one week. Furthermore, it is vital for the sensor&#8217;s functionality to place the microsphere at the tip&#8217;s center as lateral deviation in microsphere attachment easily results in inconsistent measurements.
Placing the cantilever on the glass block and fixing it with a fitting spring is a delicate and error-prone process that needs meticulous attention and steady hands. Because cantilevers are very likely to be destroyed by untrained operators, we recommend conducting several test runs and practice to comfortably handle the AFM&#8217;s easily breakable components.
A clean glass block is imperative to properly calibrate the device and obtain reliable measurements. Dirt or dust on the block&#8217;s optical surface may prevent proper laser alignment on the photodetector. Therefore, if problems during the laser alignment are encountered, washing off the glass block with ethanol a second time might be necessary.
Data analysis or determination of the elastic modulus can be performed using the Hertz model as described previously11,12. In short, the data generated by indentation are plots of force over movement of the cantilever tip. During the measurements, the cantilever is moved in the direction of the sample. This leads to the cantilever making contact with the sample and subsequently to its bending in the direction opposite to the one in which it originally moved. Simultaneously, an indentation of the sample by a certain amount occurs. In order to use the Hertz-fit model, the sample&#8217;s indentation has to be calculated and adjusted to isolate the cantilever bending parameter. The parameter describing the sample is Poisson&#8217;s ratio, which depends on the material investigated. For soft biological samples, Poisson&#8217;s ratio is often set to 0.515. As mentioned above, the shape of the used indenter is relevant to the calculation of Young&#8217;s modulus, as it dictates the extensions that have to be made to the original Hertz equation. In case of the described experiment, a spherical indenter shape is assumed due to the usage of microspheres.
While AFM may offer new and interesting possibilities of gathering data, the consistency and reliability of the yielded data strongly depend on the experience of the respective operator. Several of the steps outlined above are prone to human error and require patience and meticulousness to execute them properly.
Due to the many sensitive variables that can affect measurement results, the absolute force values reported in this study cannot be generalized but are rather specific for our experimental setup. When using this technique to evaluate the tissue degeneration of cartilage, some normalizing measurements on different spatial patterns first need to be performed to scale the results to the specific experimental measurement settings present. The relationship of the different elasticity moduli and the spatial patterns will, however, not be affected.

Articular cartilage is an avascular, aneural tissue. Sparsely scattered chondrocytes produce, organize, and maintain an expansive extracellular matrix (ECM) into which they are embedded. As a distinct and specialized part of the ECM, chondrocytes are surrounded by a thin layer of specialized matrix known as the pericellular matrix (PCM). The PCM acts as a mechanosensitive cell-matrix interface1 that protects the chondrocytes2 and modulates their biosynthetic response3. As previously described4, in healthy cartilage, chondrocytes are arranged in specific, distinct spatial patterns that are specific for each tissue layer and joint4,5 and depend on joint-specific mechanical loading mechanisms6. These patterns change from pairs and strings in healthy cartilage to double strings with the onset of osteoarthritis (OA). With further progression of the disease the chondrocytes form small clusters, increasing gradually in size to big clusters in advanced OA. A complete loss of any organizational structure and induction of apoptosis is observed in end stage OA. Thus, chondrocyte cellular arrangement can be used as an image-based biomarker for OA progression4.
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 OA. Atomic force microscopy (AFM) has emerged as a powerful tool to qualitatively and quantitatively characterize the biomechanical properties of specific biological target structures on a microscopic scale, measuring a wide range of force, from piconewton to the micronewton. The major application of AFM is to measure the surface topography and mechanical properties of samples at subnanometer resolution7. The measurement device consists of three main components: 1) An AFM probe, which is a sharp tip mounted on a cantilever and is used for the direct interaction with the surface of the sample. When force is applied to the cantilever, deformation of the latter occurs according to the measured tissue&#39;s properties. 2) An optical system that projects a laser beam onto the cantilever, which is then reflected to a detector unit. 3) A photodiode detector that catches the light deflected from the cantilever. It converts the received information regarding the laser deflection by the cantilever into a force curve that can be analyzed.
Thus, the main principle of AFM is the detection of the force acting between the AFM probe and the target structure of the sample. The force curves obtained describe the mechanical properties of the target structures on the sample surface like elasticity, charge distribution, magnetization, yield stress, and elastic plastic deformation dynamics8. An important advantage of AFM over other imaging techniques is that AFM can be used to measure the mechanical properties of live cells in medium or tissues in a native state without damaging the tissue. AFM can operate both in liquid or dry conditions. There is no requirement for sample preparation. AFM provides the possibility to image a specimen and measure its mechanical properties simultaneously in specimens that are near physiological conditions. In the present study we describe a novel approach to assess OA progression by measuring the elasticity of the PCM and ECM in native articular cartilage. The correlation of spatial organization of chondrocytes with the degree of local tissue degeneration provides a completely new perspective for early detection of OA. The functional relevance of these patterns has not been evaluated so far, however. Because the major function of articular cartilage is load bearing at low friction, the tissue must possess elastic properties. AFM allows measuring not only the elasticity of the ECM but also of the spatial cellular patterns embedded into their PCM. The observed correlation of elasticity with spatial pattern change of the chondrocytes is so strong that measuring elasticity alone may allow stratification of local tissue degeneration.
Elastic moduli of the PCM and ECM were assessed in 35 &#181;m-thin sections using an AFM system integrated into an inverted phase contrast microscope that allowed simultaneous visualization of the cartilage sample. This protocol is based on a study already published from our laboratory9 and specifically describes how to characterize the spatial arrangement of the chondrocytes and how to measure the elasticity of their associated PCM and ECM. With each pattern change of the chondrocytes, significant changes in elasticity can also be observed for both the PCM and ECM, allowing this technique to be used to directly measure the stage of degeneration of the cartilage.
This validated approach opens up a new way to evaluate OA progression and therapeutic effects at early stages before macroscopic tissue degradation actually starts to appear. Performing AFM measurements consistently is an arduous process. In the following protocol we describe how to prepare the sample to be measured by AFM, how to perform the actual AFM measurements starting with preparation of the cantilever, how to calibrate the AFM, and then how to perform the measurements. Step-by-step instructions give a clear and concise approach to obtain reliable data and provide basic strategies for processing and interpreting it. The discussion section also describes the most common pitfalls of this rigorous method and provides helpful troubleshooting tips.

<table>
	<tbody>
		<tr>
			<td>Amphotericin B</td>
			<td>Merck</td>
			<td>A2942</td>
			<td></td>
		</tr>
		<tr>
			<td>Atomic Force Microscope (AFM)</td>
			<td>CellHesion 200, JPK Instruments, Berlin, Germany</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM head</td>
			<td>(CellHesion 200) JPK</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>Biocompatible sample glue</td>
			<td>JPK Instruments AG, Berlin, Germany</td>
			<td>H000033</td>
			<td></td>
		</tr>
		<tr>
			<td>Cantilever</td>
			<td>tip C, k &#188; 7.4 N/m, All-In-One-AleTl, Budget Sensors, Sofia, Bulgaria</td>
			<td>AIO-TL-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified Eagle&#39;s medium (DMEM)</td>
			<td>Gibco, Life Technologies, Darmstadt, Germany</td>
			<td>41966052</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted phase contrast microscope (Integrated with 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</td>
			<td>(Merck KGaA, Darmstadt, Germany)</td>
			<td>F1315</td>
			<td></td>
		</tr>
		<tr>
			<td>Microspheres</td>
			<td>Polysciences</td>
			<td>07313-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Sigma</td>
			<td>P4333</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish heater associated with AFM</td>
			<td>JPK Instruments AG, Berlin, Germany</td>
			<td>T-05-0117</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Feather</td>
			<td>2023-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue culture dishes</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></td>
		</tr>
	</tbody>
</table>

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.

Along the physiopathological model from strings to double strings, to small and finally to big clusters, both ECM (Figure 3A) and PCM (Figure 3B) elastic moduli decreased significantly between each pattern change. The only exception was the difference in ECM between strings and double strings (p = 0.072). The results show that the ECM/PCM ratio (Figure 4B) did not change significantly, whereas a marked decrease in the absolute differences in elasticity between ECM and PCM was observed (Figure 4A). Furthermore, the results do not show any significant association concerning the ECM/PCM ratio or associated cellular spatial changes (r = -0.099, p = 0.281).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61041/61041fig01.jpg" /> 
Figure 1: Schematic representation of a force-distance curve in the AFM contact mode. As the probe approaches the surface, the forces are too small to give a measurable deflection of the tip at first, thus leaving the tip in its undisturbed position (1). Then, when the cantilever is very close to the sample, due to adhesive forces active between the tip and the probe, the cantilever actually quickly snaps towards the sample (2). With the probe further approaching the sample, the repulsive deflection then faces against the movement of direction, with an almost linear function of height and deflection until the vertical deflection reaches the relative set point value (3). When retracting (4), in addition to the lowering deflecting forces, adhesion forces are also present while the cantilever is retracted in the Z-axis from the sample. As the AFM probe is pulled off the contact with the sample, it first gets &#8220;stuck&#8221; before it is able to loosen from the adhesion at the interface, even leading to a short negative deflection of the cantilever, before again reaching its unbent neutral position without contact with the probe (5). The extent of deflection is expressed in the force working on the cantilever expressed in nanonewtons. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61041/61041fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61041/61041fig02.jpg" /> 
Figure 2: Representative spatial characterization of chondrocytes and AFM measurements of the extracellular matrix (ECM) and pericellular matrix (PCM). (A-D) Characterization of the cellular patterns: strings (A), double strings (B), small clusters (C), and big clusters (D). The elastic moduli of the PCM (red circles) and ECM (blue area) (E/F) were assessed for the different cellular patterns in osteoarthritic cartilage. Measurement sites for the ECM and PCM were selected by the experimenter and are graphically indicated by black crosses. The cantilever tip used for the measurements is marked by a white star. Scale bars represent 10 &#181;m (A-D), and 100 &#181;m (E/F). The figure is adapted and modified from Danalache et al.9. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61041/61041fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61041/61041fig03.jpg" /> 
Figure 3: Comparison of the quantified Young&#8217;s moduli of the extracellular matrix (ECM) and the pericellular matrix (PCM) as a function of spatial chondrocyte organization. With progressive pathological spatial chondrocyte organization, a gradual decrease of elasticity was noted in the boxplots for both the ECM (A) and the PCM (B) (*p &#60; 0.05, ***p &#60; 0.001). Abbreviations: SS: single strings, DS: double strings, SC: small clusters, BC: big clusters. The figures are taken from Danalache et al.9. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61041/61041fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61041/61041fig04.jpg" /> 
Figure 4: Relationship of the Young&#8217;s moduli of the extracellular matrix (ECM) and the pericellular matrix (PCM) as a function of cellular spatial organization. An increasingly pathological spatial chondrocyte organization was associated with a decrease of the Young&#8217;s moduli for both ECM and PCM (A). While these spatial changes took place, the ratio of ECM and PCM elasticity remained constant, showing no significant changes (B). The data are presented as a line diagram with mean &#177; standard error (A) and boxplots (B). Abbreviations: SS: single strings, DS: double strings, SC: small clusters, BC: big clusters. The figures are taken from Danalache et al.9. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61041/61041fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1: Representative force curve obtained by indentation of the pericellular matrix (PCM) of a single string pattern showing the fit results (orange arrow) as well as the residual root mean square (residual RMS; black arrow). The fit results include the contact point between sample and tip, the Young&#8217;s Modulus, and the baseline. The residual RMS displayed below describes the difference between the fit and the force data, thereby representing the quality of a force curve fit. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61041/61041supfig01.jpg" target="_blank">Please click here to download this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Table 1. Parameters for gluing a microsphere on the AFM-probe</td>
		</tr>
		<tr>
			<td>Parameters</td>
			<td>Value</td>
		</tr>
		<tr>
			<td>Setpoint</td>
			<td>5.0 V</td>
		</tr>
		<tr>
			<td>Adjust baseline</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Pulling length</td>
			<td>90.0 &#181;m</td>
		</tr>
		<tr>
			<td>Z movement</td>
			<td>Constant speed</td>
		</tr>
		<tr>
			<td>Extend speed</td>
			<td>5.0 &#181;m/s</td>
		</tr>
		<tr>
			<td>Extend time</td>
			<td>18.0 s</td>
		</tr>
		<tr>
			<td>Contact time</td>
			<td>90.0 s</td>
		</tr>
		<tr>
			<td>Delay mode</td>
			<td>Constant force</td>
		</tr>
		<tr>
			<td>Sample rate</td>
			<td>2000 Hz</td>
		</tr>
	</tbody>
</table>
Table 1. Parameters for gluing a microsphere on the AFM-probe.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Table 2. Approach parameters</td>
		</tr>
		<tr>
			<td>Approach Parameters</td>
			<td>Value</td>
		</tr>
		<tr>
			<td>Approach IGain</td>
			<td>5.0 Hz</td>
		</tr>
		<tr>
			<td>Approach PGain</td>
			<td>0.0002</td>
		</tr>
		<tr>
			<td>Approach target height</td>
			<td>10.0 &#181;m</td>
		</tr>
		<tr>
			<td>Approach setpoint</td>
			<td>5.00 V</td>
		</tr>
		<tr>
			<td>Approach baseline</td>
			<td>0.00 V</td>
		</tr>
	</tbody>
</table>
Table 2. Approach parameters.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Table 3. Run parameters</td>
		</tr>
		<tr>
			<td>Parameters</td>
			<td>Value</td>
		</tr>
		<tr>
			<td>Setpoint</td>
			<td>1.0 V</td>
		</tr>
		<tr>
			<td>Adjust baseline</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Pulling length</td>
			<td>90.0 &#181;m</td>
		</tr>
		<tr>
			<td>Z movement</td>
			<td>Constant speed</td>
		</tr>
		<tr>
			<td>Extend speed</td>
			<td>5.0 &#181;m/s</td>
		</tr>
		<tr>
			<td>Extend time</td>
			<td>18.0 s</td>
		</tr>
		<tr>
			<td>Contact time</td>
			<td>0.0 s</td>
		</tr>
		<tr>
			<td>Delay mode</td>
			<td>Constant force</td>
		</tr>
		<tr>
			<td>Sample rate</td>
			<td>2000 Hz</td>
		</tr>
	</tbody>
</table>
Table 3. Run parameters.

Watch this Scientific Journal Video about Application of Atomic Force Microscopy to Detect Early Osteoarthritis at JoVE.com

Application of Atomic Force Microscopy to Detect Early Osteoarthritis

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 ...

application-of-atomic-force-microscopy-to-detect-early-osteoarthritis

Nanyang Technological University

Research

JoVE Journal

Bioengineering

9.0K Views.  University Hospital of Tübingen. We present a method to investigate early osteoarthritic changes at the cellular level in articular cartilage by using atomic force microscopy (AFM).

Video: Application of Atomic Force Microscopy to Detect Early Osteoarthritis

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, ...

Shock Wave Application to Cell Cultures

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to target cells or tissue without any subsequent damage. Therefore, in vitro experiments are of increasing interest. Various methods of applying shock waves onto cell cultures have been described. In general, all existing models focus on how to best apply shock waves onto cells.
However, this question remains: What happens to the waves after passing the cell culture? The difference of the acoustic impedance of the cell culture medium and the ambient air is that high, that more than 99% of shock waves get reflected! We therefore developed a model that mainly consists of a Plexiglas built container that allows the waves to propagate in water after passing the cell culture. This avoids cavitation effects as well as reflection of the waves that would otherwise disturb upcoming ones. With this model we are able to mimic in vivo conditions and thereby gain more and more knowledge about how the physical stimulus of shock waves gets translated into a biological cell signal (&#8220;mechanotransduction&#34;).

Shock waves nowadays are well known for their regenerative effects. Therefore in vitro experiments are of increasing interest. We therefore developed a model for in vitro shock wave trials (IVSWT) that enables us to mimic in vivo conditions thereby avoiding distracting physical effects.

Shock waves nowadays are well known for their regenerative effects. Basic research findings showed that shock waves do cause a biological stimulus to ...

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. Kelvin probe force microscopy (KPFM) is a module of atomic force microscopy (AFM) that measures the contact potential difference between surfaces at the nano-scale. The combination of KPFM with AFM allows for the simultaneous generation of surface potential and topographical maps of biological samples such as bacterial cells. Here, we employ KPFM to examine the effects of surface potential on microbial adhesion to medically relevant surfaces such as stainless steel and gold. Surface potential maps revealed differences in surface potential for microbial membranes on different material substrates. A step-height graph was generated to show the difference in surface potential at a boundary area between the substrate surface and microorganisms. Changes in cellular membrane surface potential have been linked with changes in cellular metabolism and motility. Therefore, KPFM represents a powerful tool that can be utilized to examine the changes of microbial membrane surface potential upon adhesion to various substrate surfaces. In this study, we demonstrate the procedure to characterize the surface potential of individual methicillin-resistant Staphylococcus aureus USA100 cells on stainless steel and gold using KPFM.

Here, we present a protocol explaining the use of Kelvin probe force microscopy as a tool for generating high resolution nano-scale surface potential maps. This tool was applied to assess the role of surface potential on the binding capacity of microorganisms to substrate surfaces.

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. ...

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 ...

High Throughput Traction Force Microscopy Using PDMS Reveals Dose-Dependent Effects of Transforming Growth Factor-&#946; on the Epithelial-to-Mesenchymal Transition

Cellular contractility is essential in diverse aspects of biology, driving processes that range from motility and division, to tissue contraction and mechanical stability, and represents a core element of multi-cellular animal life. In adherent cells, acto-myosin contraction is seen in traction forces that cells exert on their substrate. Dysregulation of cellular contractility appears in a myriad of pathologies, making contractility a promising target in diverse diagnostic approaches using biophysics as a metric. Moreover, novel therapeutic strategies can be based on correcting the apparent malfunction of cell contractility. These applications,&#160;however, require direct quantification of these forces.
We have developed silicone elastomer-based traction force microscopy (TFM) in a parallelized multi-well format. Our use of a silicone rubber, specifically polydimethylsiloxane (PDMS), rather than the commonly employed hydrogel polyacrylamide (PAA) enables us to make robust and inert substrates with indefinite shelf-lives requiring no specialized storage conditions. Unlike pillar-PDMS based approaches that have a modulus in the GPa range, the PDMS used here is very compliant, ranging from approximately 0.4 kPa to 100 kPa. We create a high-throughput platform for TFM by partitioning these large monolithic substrates spatially into biochemically independent wells, creating a multi-well platform for traction force screening that is compatible with existing multi-well systems.
In this manuscript, we use this multi-well traction force system to examine the Epithelial to Mesenchymal Transition (EMT); we induce EMT in NMuMG cells by exposing them to TGF-&#946;, and to quantify the biophysical changes during EMT. We measure the contractility as a function of concentration and duration of TGF-&#946; exposure. Our findings here demonstrate the utility of parallelized TFM in the context of disease biophysics.

We present a high throughput traction force assay fabricated with silicone rubber (PDMS). This novel assay is suitable for studying physical changes in cell contractility during various biological and biomedical processes and diseases. We demonstrate this method's utility by measuring a TGF-&#946; dependent increase in contractility during the epithelial-to-mesenchymal transition.

Cellular contractility is essential in diverse aspects of biology, driving processes that range from motility and division, to tissue contraction and ...

Fabrication and Implementation of a Reference-Free Traction Force Microscopy Platform

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their microenvironment. While many approaches exist for measuring cell-induced material strain, here we provide a methodology for monitoring strain with sub-micron resolution in a reference-free manner. Using a two-photon activated photolithographic patterning process, we demonstrate how to generate mechanically and bio-actively tunable synthetic substrates containing embedded arrays of fluorescent fiducial markers to easily measure three-dimensional (3D) material deformation profiles in response to surface tractions. Using these substrates, cell tension profiles can be mapped using a single 3D image stack of a cell of interest. Our goal with this methodology is to make traction force microscopy a more accessible and easier to implement tool for researchers studying cellular mechanotransduction processes, especially newcomers to the field.

This protocol provides instructions for implementing multiphoton lithography to fabricate three-dimensional arrays of fluorescent fiducial markers embedded in poly(ethylene glycol)-based hydrogels for use as reference-free, traction force microscopy platforms. Using these instructions, measurement of 3D material strain and calculation of cellular tractions is simplified to promote high-throughput traction force measurements.

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their ...