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

Summary

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.

Abstract

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.

Introduction

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 scales^1,2. 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 systems^3,4,5,6. 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 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 damage^15,16,17. 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 arthroscopy^18,19,20.

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 others^23,24. 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 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'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.

Protocol

Femoral condyles collected from patients undergoing total knee arthroplasty at the University Hospital of Tübingen, 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

1. Tissue preparation
   1. Following post-operative resection, place the cartilage samples in a container filled with Dulbecco's modified Eagle'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.
   2. Cut the cartilage away from the bone using a scalpel.
2. Cartilage disc generation
   1. 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.
   2. 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's topmost layer (the superficial zone of the articular cartilage) does not face the blade
   3. 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.
   4. 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.
3. 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.
   1. 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.
   2. Using a standard cryotome, section the tissue sideways at a 60 µm 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.
   3. Collect the sections on a glass slide and remove the water-soluble embedding medium by washing three times with phosphate-buffered saline (PBS).

2. Cartilage disc sorting as a function of the cellular spatial pattern

1. Staining of the disc-shaped cartilage samples
   1. Place one cartilage disc (section 1.2) in each well of a 96-well plate and add 130 µL of cell permeable fluorescence dye at a dilution of 1:1,000 to each well.
   2. 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 °C.
2. Staining of the 60 µm cartilage slices
   1. Gently place the cartilage disc sections (section 1.3) on glass microscope slides with the help of forceps.
   2. Cover the cartilage sections with mounting medium containing nuclear DAPI counterstaining and gently place coverslips that are suitable for fluorescence microscopy.
   3. Seal the edges of each coverslip with regular transparent nail polish and allow to dry for 3 min.
3. 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).
   1. Place the 96-well plate on the plate holder of the fluorescence microscope.
   2. 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.
   3. 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 disc^27,28.
4. Image acquisition of the entire cartilage discs
   1. 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.
   2. 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.
   3. 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.
   4. Adjust the exposure/light intensity so that the cells can be clearly visualized from the background. At this point, the picture'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's edge often emit a higher fluorescence signal than the cells in the center, the exposure/light intensity settings must be adapted.
      1. 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's image software, visualize all the cells residing within the disc.
   5. Select the software's Focus Map Point option, and then select each individual tile by left-clicking in the center of it.
   6. 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.
   7. 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.
      1. 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.
   8. Save, export, and correctly annotate the images.

3. Biomechanical approach of cartilage explants

1. Sample preparation for AFM measurements
   1. 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.
   2. Cover the discs with 2.5 mL of Leibovitz'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.
2. Loading of the samples into the AFM
   1. Position the Petri dish in the AFM device's sample holder and turn on the Petri dish heater set to 37 °C. Allow the tissue culture dish to reach the desired temperature. This is done to exclude possible artifacts caused by temperature variation.
3. AFM-cantilever calibration
   1. Initialize the software setup as previously described by Danalache et al.²⁹.
   2. 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's reflective surface is straight and parallel to the AFM holder.
   3. 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.
   4. Carefully place a silicone skirt (silicone membrane) on the base of the cantilever holder in order to prevent medium condensation in the AFM head.
   5. Lower the cantilever in 100 µm steps using the stepper motor function until it is completely submerged in the medium.
   6. Run a scanner approach with the approach parameters described by Danalache et al.²⁹. Retract the cantilever by 100 µm once the bottom of the Petri dish is reached.
   7. 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.
   8. Using the stepper motor function, retract the cantilever to 1,000 µm.
4. 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.
   1. 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.
   2. 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.
   3. 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.
   4. Switch from brightfield to fluorescence view and visually identify the top of the disc.
   5. 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.
   6. Run a scanner approach and, once the surface of the cartilage disc is reached, retract the cantilever by 100 µm.
5. Force-distance curve measurements
   1. 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.
   2. Acquire five force-distance curves on each measurement site. Retract the cantilever by 500 µm 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.
   3. Inspect the force-distance curves and save them.
6. Estimation of Young's moduli using the Hertz fit model
   1. 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.
   2. Select the Hertz fit model and then select the Elasticity fit option.
      1. 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.
   3. 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 µm. We recommend fitting the force-distance curve until the maximum applied force is reached (setpoint).
   4. Check visually the force-distance curve fit to ensure correctness. This step has to be done for each of the force-distance curves analyzed.
7. 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.
   1. Open each of the generated force-distance curves in the data analysis software and select the Hertz fit Model as the analysis process.
   2. Apply the Subtract Baseline Offset option to zero the vertical deflection axis (y-axis) and select the Offset + Tilt function.
   3. Use the Find Contact Point function to automatically identify the contact point, which is automatically brought to an x-coordinate of zero.
   4. Subtract the distance accounting solely for cantilever deflection from the raw piezo height during the indentation using the Vertical Tip Position function.
   5. 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).
   6. Read and document the indentation from the X Min box in the parameter tab. Save and document the results.

4. Statistical analysis

1. Open the statistical software. Select New Dataset from the drop-down menu.
2. 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's moduli.
3. In the data view tab, enter the measured Young'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.
4. Select Young'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.
5. To conduct a statistical analysis, choose Dependent Samples in the nonparametric test section of the analyze menu bar tab. Select Young'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.
6. Incorporate the nonparametric test'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.

Results

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 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 ...

Discussion

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 cartilage^27,31. The earliest pathological events occurring in OA are proteoglycan depletion coupled with collagen network disruption^32,33

Materials

Name	Company	Catalog Number	Comments
Amphotericin B	Merck KGaA, Darmstadt, Germany	1397-89-3
Atomic force microscop (AFM) head	CellHesion 200, Bruker Nano GmbH, Berlin, Germany	JPK00518
Biocompatible sample glue	Bruker Nano GmbH, Berlin, Germany	H000033
Calcein AM	Cayman, Ann Arbor, Michigan, USA	14948	Cell membrane permeable stain, used for cartilage disc sorting- top view imaging
Cantilever	Bruker Nano GmbH, Berlin, Germany	SAA-SPH-5UM	Frequency Nom: 30KHz, k: 0.2N/m, lenght nom: 115μm, width nom: 40μm,  geometry: rectangular, cylindrical tip with a 5μm end radius
Cartilage ctting device	Self-made	n/a	Cutting plastic device containing predefined wholes of 4mmx1mm
CDD camera integrated in the AFM	The Imaging Source Europe GmbH, Bremen, Germany	DFK 31BF03
CDD camera integrated in the fluorescence microscope	Leica Biosystems, Wetzlar, Germany	DFC3000G
Cryotome	Leica Biosystems, Wetzlar, Germany	CM3050S
Data Processing Software for the AFM	Bruker Nano GmbH, Berlin, Germany	n/a	Version 5.0.86,  can be downloaded for free from the following website https://customers.jpk.com
Dulbecco's modified Eagle's medium (DMEM)	Gibco, Life Technologies, Darmstadt, Germany	41966052
Fluorescence Microscope (Leica DMi8)	Leica Biosystems, Wetzlar, Germany	11889113
Glass block cantiliver holder	Bruker Nano GmbH, Berlin, Germany	SP-90-05	Extra long glass block with angled faces, designed especially for the use with the JPK PetriDishHeaterTM (Bruker).
Inverted phase contrast microscope (integrated in the AFM)	AxioObserver D1, Carl Zeiss Microscopy, Jena, Germany	L201306_03
Leibovitz's L-15 medium without L-glutamine	Merck KGaA, Darmstadt, Germany	F1315
Microscope glass slides	Sigma-Aldrich, St. Louis, Missouri, USA	CLS294775X50
Mounting medium With DAPI	ibidi GmbH, Gräfelfing, Germany	50011	Mounting media with nuclear DAPI (4′,6-diamidino-2-phenylindole) counterstaining used for cartilage discs  side view imaging
Penicillin-Streptomycin	Sigma-Aldrich, St. Louis, Missouri, USA	P4333
Petri dish heater associated with AFM (Petri Dish Heater)	Bruker Nano GmbH, Berlin, Germany	T-05-0117
Scalpel	Feather Medical Products, Osaka, Japan	2023-01
Silicone Skirt	Bruker Nano GmbH, Berlin, Germany	n/a	Protective silicone membrane (D55x0.25) which is placed on the basis of the base of the glas block to prevent  medium condensation in the AFM head.
Statistical program - SPSS	IBM, Armonk, New York, USA	SPSS Statistics 22	Vesion 280.0.0.0 (190)
Tissue culture dishes	TPP Techno Plastic Products AG, Trasadingen, Switzerland	TPP93040
Tissue-tek O.C.T. Compound	Sakura Finetek, Alphen aan den Rijn, Netherlands	SA6255012	Water-soluble embedding medium