The authors would like to thank Dr. Richard Waugh and Luis Delgadillo for the generous use of their pH meter and osmometer. Additionally, the authors would like to thank Andrea Lee for contributing to the initial development of the mechanical testing system. This study was funded by NIH P30 AR069655.

All animal work was approved by the University of Rochester Committee on Animal Resources.
1. Solutions
<ol>
	<li>Prepare Hank&#39;s Balanced Salt Solution (1X HBSS) containing calcium, magnesium and no phenol red. Adjust the pH to 7.4 by adding small amounts of HCl or NaOH.</li>
	<li>Adjust the osmolarity to 303 mOsm by adding NaCl or deionized water. Use the buffer during the dissections, specimen preparation and mechanical testing.</li>
</ol>
2. Dissections of the Distal Femur and Proximal Humerus with Fully Intact Articular Cartilage
<ol>
	<li>Euthanize the mouse according to institutional guidelines using a CO2 inhalation chamber followed by cervical dislocation. Follow aseptic technique throughout. 
	NOTE: Mice between 8 and 81 weeks old of any strain and sex may be used.</li>
	<li>Use the following surgical tools for dissections: micro-scissors (used for making incisions), standard dissecting scissors (used for cutting the bone), scalpel with #11 scalpel blade (used for cutting soft tissue), jeweler&#39;s forceps (used for removal of soft tissue) and standard forceps (used for peeling the soft tissue and the skin).</li>
	<li>For dissections of the distal femur follow the instructions below.
	<ol>
		<li>Position the mouse in a supine position.</li>
		<li>Make a small (~5 mm) incision in the skin on the anterior portion of the knee.</li>
		<li>Extend the incision all the way around the knee and pull back the skin to expose the knee joint and leg muscles.</li>
		<li>Starting from the proximal end of the femur, use a scalpel blade (#11) to cut along the distal direction. Position the blade between the quadriceps muscle-tendon unit and the anterior side of the femoral shaft. Extend the cut towards and past the patella and finish by cutting through the middle of the patellar tendon, thereby removing the quadriceps muscle-tendon unit.</li>
		<li>Starting from the proximal end of the femur, use a scalpel blade (#11) to cut along the distal direction. Position the blade between the hamstring muscle-tendon unit and the posterior side of the femoral shaft. Once the cut approaches the knee joint, begin cutting through the soft tissue, avoiding contact between the scalpel and the articular surface on the distal condyles of the femur. Finish the cut past the knee joint.</li>
		<li>Pull back the quadriceps and hamstring muscles to expose the femur. Cut away any excess muscle on the lateral and medial sides of the femur.</li>
		<li>Cut the calf muscle on the posterior side of the proximal tibia and flip the leg to visualize the posterior side of the femur.</li>
		<li>Expose the distal condyles of the femur and posterior surface of the proximal tibia by removing excess tissue around the knee joint.</li>
		<li>Cut the anterior and posterior cruciate ligaments using a scalpel, cutting away from the femoral condyles. Pull the tibia away from the femur and cut all the ligaments to separate the lower leg from the femur.</li>
		<li>Using standard dissecting scissors, cut through the femur at the proximal end of the bone (8 mm above the tibio-femoral joint) from the lateral side. After cutting the bone, wipe away any visible marrow on the outside of the bone to avoid possible contamination from bone marrow cells. 
		NOTE: Cutting from lateral side reduces the risk of crack propagation along the bone.</li>
		<li>Remove surrounding soft tissues (i.e., ligaments and excess muscle) from the femur using jeweler&#39;s forceps and expose the cartilage on both condyles at the distal end of the femur. Avoid contact between the cartilage and forceps.</li>
	</ol>
	</li>
	<li>For dissections of the humerus follow the instructions below.
	<ol>
		<li>Position the mouse in a supine position.</li>
		<li>Make an incision (~5 mm) in the skin on the posterior side of the elbow using micro-scissors, extend the incision around the elbow and pull back the skin to expose the muscles of the arm and shoulder.</li>
		<li>Starting from the proximal end of the humerus, use a scalpel blade (#11) to cut along the distal direction. Position the blade between the triceps muscle-tendon unit and the posterior side of the humerus. Extend the cut towards the distal end of the humerus and finish by cutting through the triceps tendon.</li>
		<li>Then pull back the triceps towards the proximal end of the humerus until the humeral head is exposed.</li>
		<li>Cut the connective tissue around the humeral head using a scalpel blade (#11) without touching the articular surface and remove the limb (arm and shoulder) from the body. Periodically hydrate the articular surface of the humeral head with HBSS.</li>
		<li>Disconnect the humerus from the arm by first breaking off the proximal end of the ulna on the posterior side of the arm using forceps and then cutting the connective tissue around the distal end of the humerus. Remove any excess tissue on the humerus.</li>
		<li>Cut off the deltoid tuberosity on the posterior side of the humerus using standard dissecting scissors.</li>
	</ol>
	</li>
	<li>Place the dissected specimen(s) into a 1.5 mL microcentrifuge tube containing HBSS buffer.</li>
</ol>
3. Live (Calcein AM) / Dead (Propidium Iodide) Staining Protocol
<ol>
	<li>Stain using calcein AM as follows.
	<ol>
		<li>Make a stock solution of calcein AM by adding 12.5 &#181;L of dimethyl sulfoxide (DMSO) into a vial of 50 &#181;g of calcein AM, resulting in a stock concentration of 4 mg/mL (4.02 mM).</li>
		<li>Dilute the stock calcein solution 1:400 in HBSS to reach a concentration of 10 &#181;g/mL (10.05 &#181;M) of calcium AM (for example, add 1.25 &#181;L of stock solution to 500 &#181;L of HBSS).</li>
		<li>Centrifuge the diluted staining solution for 5 s at 2,000 x g to ensure that all of the dye, which may occasionally stick to the walls of the tube, mixes with the buffer. Do not remove the supernatant. Vortex the solution to ensure proper mixing.</li>
		<li>Put the dissected specimen into a 1.5 mL microcentrifuge tube containing 500 &#181;L of the diluted staining solution. Incubate the specimen at 37 &#176;C for 30 min in a thermomixer while agitating at 800 rpm. Make sure the tubes are covered in aluminum foil to protect from exposure to light.</li>
		<li>Transfer the specimen into calcein AM-free HBSS buffer. At this point, the specimen is ready for mechanical testing.</li>
	</ol>
	</li>
	<li>Prepare propidium iodide (PI) staining solution as follows.
	<ol>
		<li>Prepare the PI staining solution by diluting the purchased stock solution (1 mg/mL, 1.5 mM) 1:25 in HBSS to reach 40 &#181;g/mL (60 &#181;M) of PI. For example, add 40 &#181;L of stock solution to 1000 &#181;L of HBSS.</li>
		<li>Keep the PI staining solution covered in aluminum foil to protect it from exposure to light. Use the staining solution immediate after the mechanical test to detect injured cells (see section 4). 
		NOTE: One can perform the PI staining prior to loading as well to more rigorously assess the quality of the dissections.</li>
	</ol>
	</li>
</ol>
4. Mechanical Testing Protocol
<ol>
	<li>Place the specimen (femur or humerus) onto the glass slide of a custom microscope-mounted mechanical testing device such that the articular surface on the posterior femoral condyles or humeral head is sitting on the glass (Figure 1). 
	NOTE: Fasten a 10 mm-long wooden applicator (diameter = 2 mm) to the distal side of the humerus using cyanoacrylate glue before placing it onto the device in order to stabilize the specimen on the flat surface. For a detailed description of the mechanical testing platform see Supplementary File 1: section 1.</li>
	<li>Hydrate the specimen with HBSS and place the device with the specimen onto a fluorescence microscope.</li>
	<li>Image the articular chondrocytes stained with calcein AM (excitation/emission wavelengths = 495/515 nm) before (baseline) application of the load under a fluorescence microscope with a 4X dry lens (NA = 0.13). Adjust the acquisition settings to optimize the image quality.</li>
	<li>Apply mechanical loading on top of the specimen such that articular cartilage is compressed against the cover glass.
	<ol>
		<li>For static loading, apply a prescribed static load (e.g., 0.5 N) on top of the specimen (femur or humerus). Hold the load for 5 min and then remove it.</li>
		<li>For impact, apply a prescribed impact energy (e.g., 1 mJ) onto the specimen by dropping a cylindrical impactor of known weight (e.g., 0.1 N) from a prescribed height (e.g., 1 cm). Release the load 5 s after the impact. 
		NOTE: The weight of the impactor (mg) and the prescribed height (h) can be converted into impact energy (E) using the following equation: E = mgh.</li>
	</ol>
	</li>
	<li>Incubate the specimen in PI staining solution for 5 min at room temperature.</li>
	<li>Image the articular chondrocytes stained with calcein AM and PI (excitation/emission wavelengths = 535/617 nm) under a fluorescence microscope (Figure 2).</li>
</ol>
5. Data Analysis
<ol>
	<li>Quantify the area of injured/dead cells due to the applied mechanical loading regimen.
	<ol>
		<li>Open the micrographs of the articular surface acquired before and after application of mechanical load in ImageJ18.</li>
		<li>Combine the images into a stack.</li>
		<li>Set the scale of the images based on the image resolution.</li>
		<li>Use the Polygon tool to define the area where the cells became calcein-negative (loss of green fluorescence) and PI-positive (gain of red fluorescence). These cells are considered to be injured or dead.</li>
		<li>Determine the area of injured/dead cells using the Measurement tool.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The methods described above were successfully employed to visualize viable and injured/dead in situ articular chondrocytes from mouse joints after prescribed mechanical loads or impacts. In particular, we were able to analyze the mechanical vulnerability of chondrocytes within fully intact articular cartilage from two different synovial joints: the knee joint (distal femurs) and shoulder (humeri). Our representative results show that the spatial extent of cell injury on the articular surface depends sensitively on load magnitude and impact intensity (Figure 3). Importantly, the use of this method facilitates investigations of the cellular response to mechanical loading under physiologically relevant conditions. That is, it enables testing of articular cartilage on an intact joint under physiological and supraphysiological loads (see Supplementary File 1:&#160;section 2).
Given the steep learning curve in performing dissections of murine synovial joints and challenges in preserving viable in situ chondrocytes at baseline, some protocol modification and troubleshooting may be required. The greatest risk of damaging articular chondrocytes during dissection occurs during steps 2.3.5 through 2.3.10 and 2.4.5 through 2.4.7. To minimize cell injury/death during dissections, the researcher should avoid any contact between surgical tools (e.g., the scalpel when cutting the tissue or the jeweler&#39;s forceps when removing the soft tissue) and the articular surface of the specimen. However, touching the articular surface with a glove induces little cell injury/death. In order to improve baseline cell viability, it may also be necessary to reduce the amount of soft tissue removed from the joint. Additionally, using finer tools will generally reduce dissection-induced cell death. Ultimately, to rigorously confirm the absence of dissection-associated damage of articular chondrocytes at baseline, it is advisable to stain the specimens with both permeability dyes (calcein AM and PI, prior to loading) especially while the researcher is becoming familiar with the dissection procedure (see Supplemental File 1: section 3).
Given the small size of the mouse joint and the genetic manipulability of the mouse genome, murine models provide multiple advantages over large animal models to study vulnerability of articular chondrocytes to mechanical loading. However, to the authors&#39; knowledge, no studies have previously been conducted to quantify injury/death of in situ articular chondrocytes due to mechanical loading of intact murine cartilage. Investigators typically use explants removed from joints of large animal models to investigate the extent of cell death due to mechanical injury3,4,5,6,7,8,9,10,11,12. In contrast, mouse models facilitate 1) visualization of nearly the entire articular surface on a given bone; and 2) analyses of post-loading or post-impact chondrocyte viability in intact joints without compromising native boundary conditions. Furthermore, while compressed, mouse specimens generate substantially smaller contact areas compared to large animal models; therefore, stresses are dramatically higher than in large animal models for a given load magnitude. Hence, cell injury can be induced by smaller loads. Additionally, murine models facilitate research on cartilage degeneration and, in particular, osteoarthritis, as this disease can be easily induced in mice through genetic19,20,21,22, dietary23,24 or surgical manipulations25,26,27,28. Moreover, spontaneous osteoarthritis occurs in several mouse strains including BALB/c and C57BL/629,30.
In our representative data, we used viability (live/dead) stains to quantify cell injury/death 5 min after the removal of mechanical loading. We acknowledge that these experiments may not discriminate injured cells (cells with temporarily ruptured membranes) from dead cells (cells with permanently ruptures membranes). That is, cells that are calcein negative and PI positive- indicating that the membrane was previously permeabilized-may repair their membranes over time scales ranging from seconds to several minutes31. In fact, in a separate set of experiments, we have determined that the fraction of &#34;injured&#34; cells that survive mechanical trauma is small (~5%) but significant (see Supplementary File 1: section 4). Therefore, live/dead staining is a direct measure of membrane integrity that is not always indicative of cell viability. In particular, PI-positive and calcein-negative cells are most appropriately defined as &#34;injured&#34;, where injury is defined as a (potentially temporary) loss of plasma membrane integrity due to mechanical trauma. We also acknowledge that our representative data likely reflects only immediate (necrotic) cell death. Imaging specimens at later time points (e.g., 48 h after removal of load) should enable quantification of both necrotic and apoptotic cell death.
Several limitations must be considered when using these methods. In our trial experiments for this manuscript, we used 8-10-weeks-old female BALB/c mice to demonstrate the capabilities of the testing platform (Figure 3). However, due to noticeable alterations in cell density in femurs of older mice, the assessment of load-induced cell injury becomes more challenging (though feasible) in older femurs (success rate = 40%). In contrast, no noticeable alterations in cell density on humeri were observed in 61-81-week-old C57BL/6 mice (see Supplementary File 1: section 5), thereby making the testing platform useful for ages 8-81 weeks. Another limitation is that the method was used to analyze the spatial extent of cell injury only on the articular surface of femoral condyles and humeral head. However, the method could further be extended to analyze depth-dependent spatial extent of cell injury through use of laser scanning confocal microscopy. Note that the latter method would require usage of more expensive equipment, longer image acquisition time, and more involved and time-consuming analysis. Finally, although the contact location between the articular cartilage and the cover glass was physiologically relevant for mice32,33, this location was not varied. However, our testing platform allows for variation of the contact location if the femur or humerus is gripped with a rotating armature.
In conclusion, we have developed an in vitro murine injury model that enables application of controlled mechanical loads and/or impacts onto the articular surface of intact articular cartilage. This model enables visualization of fluorescently labeled articular chondrocytes in real-time and rapid single image-based analysis of cell injury/death. Importantly, the effects of different mechanical loading regimens, different environmental conditions or different genetic manipulations on short- and/or long-term chondrocyte viability can be tested using this methodology. Thus, our platform provides a tool to interrogate basic science questions and screen therapeutic targets related to the mechanical vulnerability of chondrocytes.

Articular cartilage (AC) is a load bearing tissue that covers and protects bones in synovial joints, providing smooth joint articulation. Tissue homeostasis is dependent on the viability of chondrocytes, the sole cell type residing in AC. However, exposure of cartilage to extreme forces due to trauma (e.g., falls, vehicle accident or sports injuries) or due to post-traumatic joint instability can induce chondrocyte death, leading to irreversible breakdown of the joint (osteoarthritis)1. Furthermore, in osteochondral grafting procedures that aim to repair local defects in damaged cartilage, graft insertion-associated mechanical trauma reduces chondrocyte viability and has detrimental effects on surgical outcomes2.
Cartilage explant models are commonly used to study the susceptibility of articular chondrocytes to mechanically-induced cell death. These models typically use explants from large animals to study the effects of loading conditions, environmental conditions and other factors on cell vulnerability3,4,5,6,7,8,9,10,11,12,13,14,15. However, due to the large size of the native joints, these models generally require removal of a plug from the articular surface of an intact joint, thereby compromising native boundary conditions. Moreover, they generally require application of large mechanical loads to induce cell injury. Alternatively, murine cartilage explant models provide several advantages over larger animal models in studying the mechanical vulnerability of in situ chondrocytes. In particular, due to their smaller dimensions, these models facilitate testing of fully intact articular cartilage without altering native tissue integrity. In addition, loading of murine cartilage occurs over small contact areas such that chondrocyte death/injury can be induced with small loads (&#60;1 N). Finally, the mouse genome is easily manipulated, enabling testing of how specific genes impact the susceptibility of in situ chondrocytes to mechanical injury.
The overall goal of the method introduced in this manuscript is to quantify and visualize-in real-time-the spatial extent of in situ cell death/injury due to applied mechanical loads on fully intact mouse cartilage-on-bone explants in vitro. This method requires careful dissection of mouse synovial joints without compromising chondrocyte viability, followed by mechanical testing of vitally stained explants using a microscope-mounted device similar to a testing platform that we recently developed to quantify murine cartilage mechanical properties16. During mechanical testing, a large portion of the (intact) articular surface of the dissected bone is visible on a single fluorescence micrograph, enabling rapid analysis of cell viability after a load is applied. A similar analysis of surface cell viability in murine cartilage explants has been performed previously, but without simultaneous application of load17. Potential applications of our method include comparative studies to investigate the vulnerability of articular chondrocytes to different controlled environmental and mechanical conditions, as well as screening of treatments aimed at reducing the sensitivity of chondrocytes to mechanical loading.

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			<td>1 mg/mL solution in water, 10mL, Eugene, OR, USA</td>
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			<td>1L DMSO, anhydrous, &#8805;99.9%, St. Louis, MO, USA</td>
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			<td height="42" style="height:42px;width:223px;">HBSS (calcium, magnesium, no phenol red)&#160;</td>
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			<td>1X, 500mL, Grand Island, NY, USA</td>
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			<td height="21" style="height:21px;width:223px;">Feather surgical blade (#11)</td>
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			<td>Hatfield, PA, USA</td>
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			<td>Puteaux, France</td>
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			<td style="width:205px;">Beckman</td>
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			<td>Brea, CA, USA</td>
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			<td height="21" style="height:21px;width:223px;">Eppendorf thermomixer&#160;</td>
			<td style="width:205px;">Eppendorf AG&#160;</td>
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			<td>Hamburg, Germany</td>
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			<td height="42" style="height:42px;width:223px;">Motorized inverted research microscope</td>
			<td style="width:205px;">Olypmus</td>
			<td style="width:220px;">Model IX-81</td>
			<td>Center Valley, PA, USA</td>
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			<td height="42" style="height:42px;width:223px;">Wooden applicator</td>
			<td style="width:205px;">Puritan Medical Products Company, LLC</td>
			<td style="width:220px;">807</td>
			<td>6&#34;x100, Guilford, ME, USA</td>
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			<td height="21" style="height:21px;width:223px;">1.5 Glass coverslips</td>
			<td style="width:205px;">Warner Instruments, LLC</td>
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			<td>#1.5, 0.17mm thick, 40mm diameter, Hamden, CT, USA</td>
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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.

Six different applied loading protocols (static loading: 0.1 N, 0.5 N and 1 N for 5 min; and impact loading: 1 mJ, 2 mJ and 4 mJ) reproducibly induced quantifiable localized areas of cell injury in femoral and humeral cartilage obtained from 8-10-week-old BALB/c mice (Figure 2). Importantly, the spatial extent of chondrocyte injury on the articular surface was measured quickly and easily in ImageJ. Representative results demonstrate that the mechanical vulnerability of articular chondrocytes was affected by load magnitude and impact intensity. In particular, higher load magnitudes and higher impact intensities significantly exacerbated the spatial extent of cell injury in both femurs and humeri (Figure 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58487/58487fig1.jpg" /> 
Figure 1: Schematic representation of the custom mechanical testing device. (a) Assembled device with the cylindrical impactor used to apply prescribed mechanical loads and/or impact energies to a specimen. The device is shown without a specimen. (b) Schematic representation of the experiment. Controlled static (e.g., 0.1 N) and/or impact (e.g., 1 mJ) loading can be applied on top of the dissected specimen such that articular cartilage is compressed against the cover glass. <a href="https://www.jove.com/files/ftp_upload/58487/58487fig1large.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/58487/58487fig2.jpg" /> 
Figure 2: Representative micrograph of the articular surface after injurious mechanical loading. Representative micrograph of the articular surface on (a) the distal femoral condyles and (b) the humeral head after injurious mechanical loading (impact [1 mJ] and static loading [1 N],&#160;respectively). Green cells are vitally stained chondrocytes with intact cell membranes while red nuclei indicate injured cells with permeabilized cell membranes. (c) Zoomed-in view of the area of injured/dead cells (yellow contours) on the articular surface of the humeral head. <a href="https://www.jove.com/files/ftp_upload/58487/58487fig2large.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/58487/58487fig3v2.jpg" /> 
Figure 3: Area of injured/dead cells. Area of injured/dead cells on the articular surface of the distal femoral condyles (a, b) and the humeral head (c, d) after (a, c) static (0.1 N, 0.5 N and 1 N; n = 6 per group) and (b, d) impact (1 mJ, 2 mJ and 4 mJ; n = 6 per group) loading. All cartilage-on-bone specimens were obtained from female BALB/c 8-10 weeks old mice. Data are mean + standard deviation; brackets denote statistical significance at &#945;=0.05 determined by analysis of variance (ANOVA) test with Tukey post hoc comparisons. <a href="https://www.jove.com/files/ftp_upload/58487/58487fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1.&#160;<a href="https://www.jove.com/files/ftp_upload/58487/Supplementary_File_1.docx" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Real-time Visualization and Analysis of Chondrocyte Injury Due to Mechanical Loading in Fully Intact Murine Cartilage Explants at JoVE.com

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

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 ...

real-time-visualization-analysis-chondrocyte-injury-due-to-mechanical

Nanyang Technological University

Research

JoVE Journal

Bioengineering

6.7K Views.  University of Rochester. 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.

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

Design of a Biaxial Mechanical Loading Bioreactor for Tissue Engineering

We designed a loading device that is capable of applying uniaxial or biaxial mechanical strain to a tissue engineered biocomposites fabricated for transplantation. While the device primarily functions as a bioreactor that mimics the native mechanical strains, it is also outfitted with a load cell for providing force feedback or mechanical testing of the constructs. The device subjects engineered cartilage constructs to biaxial mechanical loading with great precision of loading dose (amplitude and frequency) and is compact enough to fit inside a standard tissue culture incubator. It loads samples directly in a tissue culture plate, and multiple plate sizes are compatible with the system. The device has been designed using components manufactured for precision-guided laser applications. Bi-axial loading is accomplished by two orthogonal stages. The stages have a 50 mm travel range and are driven independently by stepper motor actuators, controlled by a closed-loop stepper motor driver that features micro-stepping capabilities, enabling step sizes of less than 50 nm. A polysulfone loading platen is coupled to the bi-axial moving platform. Movements of the stages are controlled by Thor-labs Advanced Positioning Technology (APT) software. The stepper motor driver is used with the software to adjust load parameters of frequency and amplitude of both shear and compression independently and simultaneously. Positional feedback is provided by linear optical encoders that have a bidirectional repeatability of 0.1 &mu;m and a resolution of 20 nm, translating to a positional accuracy of less than 3 &mu;m over the full 50 mm of travel. These encoders provide the necessary position feedback to the drive electronics to ensure true nanopositioning capabilities. In order to provide the force feedback to detect contact and evaluate loading responses, a precision miniature load cell is positioned between the loading platen and the moving platform. The load cell has high accuracies of 0.15% to 0.25% full scale.

We designed a novel mechanical loading bioreactor that can apply uniaxial or biaxial mechanical strain to a cartilage biocomposite prior to transplantation into an articular cartilage defect.

We  designed a loading device that is capable of applying uniaxial or biaxial  mechanical strain to a tissue engineered biocomposites fabricated for  ...

Simultaneously Capturing Real-time Images in Two Emission Channels Using a Dual Camera Emission Splitting System: Applications to Cell Adhesion

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to investigate mechanisms governing cell adhesion under dynamic flow conditions. For instance, cancer cells labeled with multiple fluorophores can be perfused over a potentially reactive substrate to model mechanisms of cancer metastasis. However, multi-channel single camera systems and color cameras exhibit shortcomings in image acquisition for real-time live cell analysis. To overcome these limitations, we used a dual camera emission splitting system to simultaneously capture real-time image sequences of fluorescently labeled cells in the flow chamber. Dual camera emission splitting systems filter defined wavelength ranges into two monochrome CCD cameras, thereby simultaneously capturing two spatially identical but fluorophore-specific images. Subsequently, psuedocolored one-channel images are combined into a single real-time merged sequence that can reveal multiple target molecules on cells moving rapidly across a region of interest.

Dual camera emission splitting systems for two-color fluorescence microscopy generate real-time image sequences with exceptional optical and temporal resolution, a requirement of certain live cell assays including parallel plate flow chamber adhesion assays. When software is employed to merge images from simultaneously acquired emission channels, pseudocolored image sequences are produced.

Multi-color immunofluorescence microscopy to detect specific molecules in the cell membrane can be coupled with parallel plate flow chamber assays to ...

Using Cell-substrate Impedance and Live Cell Imaging to Measure Real-time Changes in Cellular Adhesion and De-adhesion Induced by Matrix Modification

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and other matrix-modifying oxidants/enzymes released during inflammation) are implicated in triggering pathological changes in cellular function, phenotype and viability in a number of diseases. Here, we describe how cell-substrate impedance and live cell imaging approaches can be readily employed to accurately quantify real-time changes in cell adhesion and de-adhesion induced by matrix modification (using endothelial cells and myeloperoxidase as a pathophysiological matrix-modifying stimulus) with high temporal resolution and in a non-invasive manner. The xCELLigence cell-substrate impedance system continuously quantifies the area of cell-matrix adhesion by measuring the electrical impedance at the cell-substrate interface in cells grown on gold microelectrode arrays. Image analysis of time-lapse differential interference contrast movies quantifies changes in the projected area of individual cells over time, representing changes in the area of cell-matrix contact. Both techniques accurately quantify rapid changes to cellular adhesion and de-adhesion processes. Cell-substrate impedance on microelectrode biosensor arrays provides a platform for robust, high-throughput measurements. Live cell imaging analyses provide additional detail regarding the nature and dynamics of the morphological changes quantified by cell-substrate impedance measurements. These complementary approaches provide valuable new insights into how myeloperoxidase-catalyzed oxidative modification of subcellular extracellular matrix components triggers rapid changes in cell adhesion, morphology and signaling in endothelial cells. These approaches are also applicable for studying cellular adhesion dynamics in response to other matrix-modifying stimuli and in related adherent cells (e.g., epithelial cells).

Here, we present a protocol to continuously quantify cell adhesion and de-adhesion processes with high temporal resolution in a non-invasive manner by cell-substrate impedance and live cell imaging analyses. These approaches reveal the dynamics of cell adhesion/de-adhesion processes triggered by matrix modification and their temporal relationship to adhesion-dependent signaling events.

Cell-matrix adhesion plays a key role in controlling cell morphology and signaling. Stimuli that disrupt cell-matrix adhesion (e.g., myeloperoxidase and ...

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 ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

A Test Bed to Examine Helmet Fit and Retention and Biomechanical Measures of Head and Neck Injury in Simulated Impact

Conventional wisdom and the language in international helmet testing and certification standards suggest that appropriate helmet fit and retention during an impact are important factors in protecting the helmet wearer from impact-induced injury. This manuscript aims to investigate impact-induced injury mechanisms in different helmet fit scenarios through analysis of simulated helmeted impacts with an anthropometric test device (ATD), an array of headform acceleration transducers and neck force/moment transducers, a dual high speed camera system, and helmet-fit force sensors developed in our research group based on Bragg gratings in optical fiber. To simulate impacts, an instrumented headform and flexible neck fall along a linear guide rail onto an anvil. The test bed allows simulation of head impact at speeds up to 8.3 m/s, onto impact surfaces that are both flat and angled. The headform is fit with a crash helmet and several fit scenarios can be simulated by making context specific adjustments to the helmet position index and/or helmet size. To quantify helmet retention, the movement of the helmet on the head is quantified using post-hoc image analysis. To quantify head and neck injury potential, biomechanical measures based on headform acceleration and neck force/moment are measured. These biomechanical measures, through comparison with established human tolerance curves, can estimate the risk of severe life threatening and/or mild diffuse brain injury and osteoligamentous neck injury. To our knowledge, the presented test-bed is the first developed specifically to assess biomechanical effects on head and neck injury relative to helmet fit and retention.

Using an anthropometric head and neck, optical fiber-based fit force transducers, an array of head acceleration and neck force/moment transducers, and a dual high speed camera system, we present a test bed to study helmet retention and effects on biomechanical measures of head and neck injury secondary to head impact.

Conventional wisdom and the language in international helmet testing and certification standards suggest that appropriate helmet fit and retention during ...

A Protocol for Real-time 3D Single Particle Tracking

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although various RT-3D-SPT methods have been put forward in recent years, tracking high speed 3D diffusing particles at low photon count rates remains a challenge. Moreover, RT-3D-SPT setups are generally complex and difficult to implement, limiting their widespread application to biological problems. This protocol presents a RT-3D-SPT system named 3D Dynamic Photon Localization Tracking (3D-DyPLoT), which can track particles with high diffusive speed (up to 20 &#181;m2/s) at low photon count rates (down to 10 kHz). 3D-DyPLoT employs a 2D electro-optic deflector (2D-EOD) and a tunable acoustic gradient (TAG) lens to drive a single focused laser spot dynamically in 3D. Combined with an optimized position estimation algorithm, 3D-DyPLoT can lock onto single particles with high tracking speed and high localization precision. Owing to the single excitation and single detection path layout, 3D-DyPLoT is robust and easy to set up. This protocol discusses how to build 3D-DyPLoT step by step. First, the optical layout is described. Next, the system is calibrated and optimized by raster scanning a 190 nm fluorescent bead with the piezoelectric nanopositioner. Finally, to demonstrate real-time 3D tracking ability, 110 nm fluorescent beads are tracked in water.

This protocol details the construction and operation of a real-time 3D single particle tracking microscope capable of tracking nanoscale fluorescent probes at high diffusive speeds and low photon count rates.

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although ...

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved oxygen concentration). Nevertheless, the morphology of cells can be a suitable indicator for optimal conditions, since it changes with dependence on the growth state, product accumulation and cell stress. Furthermore, the single-cell size distribution provides not only information about the cultivation conditions, but also about the population heterogeneity. To gain such information, a photo-optical in situ microscopy device1 was developed to enable the monitoring of the single-cell size distribution directly in the cell suspension in bioreactors. An automated image analysis is coupled to the microscopy based on a neural network model, which is trained with user-annotated images. Several parameters, which are gained from the captures of the microscope, are correlated to process relevant features of the cells, like their metabolic activity. Until now, the presented in situ microscopy probe series was applied to measure the pellet size in filamentous fungi suspensions. It was used to distinguish the single-cell size in microalgae cultivation and relate it to lipid accumulation. The shape of cellular particles was related to budding in yeast cultures. The microscopy analysis can be generally split into three steps: (i) image acquisition, (ii) particle identification, and (iii) data analysis, respectively. All steps have to be adapted to the organism, and therefore specific annotated information is required in order to achieve reliable results. The ability to monitor changes in cell morphology directly in line or on line (in a by-pass) enables real-time values for monitoring and control, in process development as well as in production scale. If the off line data correlates with the real-time data, the current tedious off line measurements with unknown influences on the cell size become needless.

In Situ Microscopy for Real-time Determination of Single-cell Morphology in Bioprocesses

A photo-optical in situ microscopy device was developed to monitor the size of single cells directly in the cell suspension. The real-time measurement is conducted by coupling the photo-optical sterilizable probe to an automated image analysis. Morphological changes appear with dependence on the growth state and cultivation conditions.

In situ monitoring in microbial bioprocesses is mostly restricted to chemical and physical properties of the medium (e.g.,pH value and the dissolved ...

Manufacturing of a Nafion-coated, Reduced Graphene Oxide/Polyaniline Chemiresistive Sensor to Monitor pH in Real-time During Microbial Fermentation

Here, we report the engineering of a solid-state micro pH sensor based on polyaniline-functionalized, electrochemically reduced graphene oxide (ERGO-PA). Electrochemically reduced graphene oxide acts as the conducting layer and polyaniline acts as a pH-sensitive layer. The pH-dependent conductivity of polyaniline occurs by doping of holes during protonation and by the dedoping of holes during deprotonation. We found that an ERGO-PA solid-state electrode was not functional as such in fermentation processes. The electrochemically active species that the bacteria produce during the fermentation process interfere with the electrode response. We successfully applied Nafion as a proton-conducting layer over ERGO-PA. The Nafion-coated electrodes (ERGO-PA-NA) show a good sensitivity of 1.71 &#937;/pH (pH 4 - 9) for chemiresistive sensor measurements. We tested the ERGO-PA-NA electrode in real-time in the fermentation of Lactococcus lactis. During the growth of L. lactis, the pH of the medium changed from pH 7.2 to pH 4.8 and the resistance of the ERGO-PA-NA solid-state electrode changed from 294.5 &#937; to 288.6 &#937; (5.9 &#937; per 2.4 pH unit). The pH response of the ERGO-PA-NA electrode compared with the response of a conventional glass-based pH electrode shows that reference-less solid-state microsensor arrays operate successfully in a microbiological fermentation.

Here, we report the protocol for the fabrication of a Nafion-coated, polyaniline-functionalized, electrochemically reduced graphene oxide chemiresistive micro pH sensor. This chemiresistor-based, solid-state micro pH sensor can detect pH changes in real-time during a Lactococcus lactis fermentation process.

Here, we report the engineering of a solid-state micro pH sensor based on polyaniline-functionalized, electrochemically reduced graphene oxide (ERGO-PA). ...