AFM analysis was performed on instrumentation supported by the Pennsylvania Muscle Institute and the Institute for Translational Medicine and Therapeutics, Perelman School of Medicine, the University of Pennsylvania. This work was supported by NIH grants HL62250 and AG047373. YHB was supported by post-doctoral fellowship from the American Heart Association.

Animal work in this study was approved by the Institutional Animal Care and Use Committees of the University of Pennsylvania. The methods were carried out in accordance with the approved guidelines.
1. Preparing the Mouse and Isolation of the Aorta
<ol>
	<li>Anesthetize a mouse with ketamine (80 - 100 mg/kg), xylazine (8 - 10 mg/kg) and acepromazine (1 - 2 mg/kg) intraperitoneally. Confirm anesthesia with a tail pinch test. Once the mouse is fully anesthetized, euthanize the mouse by cervical dislocation.</li>
	<li>Place the mouse on its back and pin the mouse to a dissection board. Clean the abdomen area with 70% (v/v) ethanol wipes.</li>
	<li>Pinch the skin at the mid-line and make a small initial incision with medium sized microsurgery scissors at the abdomen. While holding the skin with forceps, use medium sized microsurgery scissors to cut the skin and peritoneum from the abdomen to the sternum.</li>
	<li>Cut away the ribs on both sides with medium sized microsurgery scissors. Carefully remove the lungs and liver with small sized microsurgery scissors, and leave the heart and aorta. Transfer the mouse and dissection board to a dissecting microscope.</li>
	<li>Grasp the fat surrounding the aorta with small forceps and use small scissors to carefully cut away fat around the aorta.</li>
	<li>Gently grab the aorta with forceps, make one cut with small sized microsurgery scissors at the beginning of the ascending aorta and another cut at the end of the descending aorta, just above the abdominal aorta.</li>
	<li>Transfer the dissected aorta to a 60-mm dish containing 1x phosphate-buffered saline (PBS; without calcium and magnesium).</li>
	<li>Continue to dissect away any remaining fatty tissue from the aorta using small sized microsurgery scissors. Use small scissors to open the aorta longitudinally.</li>
	<li>Use small sized microsurgery scissors to cut a small piece (~ 2 x 4 mm) of the descending aorta and aortic arch for AFM analysis (Figure 1). Place the tissue in a 60-mm plastic dish and keep it moist in a drop of PBS .</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54630/54630fig1.jpg" /> 
Figure 1: An Image Showing the Location of the Different Aortic Segments in a Mouse. The aorta was isolated from the heart to the diaphragm, and a small portion of the descending aorta and the aortic arch were used to determine the elastic moduli. Scale bar, 1 mm. <a href="https://www.jove.com/files/ftp_upload/54630/54630fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Preparing Tissue Samples for the AFM Measurements
<ol>
	<li>Carefully remove the PBS using a lab-wipe without touching the aorta. Be certain that the luminal side of the tissue is facing up.</li>
	<li>Gently glue each end of the opened aorta to the plate by adding 5 - 10 &#956;l of cyanoacrylate adhesive with a gel-loading tip (Figure 2). 
	NOTE: The amount of glue needed to firmly attach the tissue at each end must be determined empirically. It is critical that the aorta is not stretched during this process.</li>
	<li>Check the glued aorta to make certain that it is not folded or floating. After air-drying the glue on the aorta (30 - 60 sec), gently add PBS and submerge the sample. 
	NOTE: Preparation the AFM (steps 3 - 5) can be carried out while the tissue is being prepared for analysis.</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54630/54630fig2.jpg" /> 
Figure 2: Cartoon of an Aortic Segment Glued onto a 60-mm Culture Dish using Cyanoacrylate Adhesive. The cyanoacrylate adhesive is being applied to the edge of an aortic sample in preparation for AFM measurements. <a href="https://www.jove.com/files/ftp_upload/54630/54630fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Loading the Probe
<ol>
	<li>Mount a fluid probe holder onto the probe load stand.</li>
	<li>Using tweezers, slide a silicon nitride AFM probe (0.06 N/m cantilever) with a spherical tip (we have used a 1-&#181;m diameter SiO2 particle but larger sizes may also be used) on the probe holder. Make sure that the AFM probe is firmly secured to the probe holder.</li>
	<li>Slide the probe holder onto the Z-scanner mount of the AFM head, and ensure that the probe holder is securely attached.</li>
</ol>
4. Aligning the Laser on the Probe
<ol>
	<li>Open the AFM software. Choose the experiment type on the software; Experiment Category to Contact Mode, Experiment Group to Contact Mode in fluid and Experiment to Contact Mode in fluid. Click Load experiment in the software; this will turn on the laser.</li>
	<li>Add 3 ml of distilled water to a 50-mm glass bottom dish and place the dish on the stage of the laser alignment unit. This unit simplifies the process of focusing the laser beam onto the back of cantilever.</li>
	<li>Mount the AFM head in an upright position onto the laser alignment unit and power on the laser alignment unit. Add approximately 50 &#956;l of distilled water to form a droplet on the AFM tip.</li>
	<li>Using the joystick, lower the AFM head so the AFM tip makes contact with water in the dish. If contact is not made between the AFM tip and water in the dish, gently raise the dish until contact is made. Adjust the focus, brightness and XY position so the AFM tip is in view on the LCD screen of the alignment unit.</li>
	<li>Position the laser spot onto the tip of the AFM probe by adjusting the laser positioning knobs of the AFM head. Using the detector positioning knobs on the AFM head, adjust the position of the photodiode until values between 0 and -1 V are obtained for the vertical deflection and ~ 0 V is obtained for the horizontal deflection.</li>
	<li>Check to ensure the laser sum signal is maximized. If necessary, repeat step 4.5 to re-adjust the position of the laser beam on the AFM tip and the position of the photodiode to obtain the maximum sum signal. 
	NOTE: It may be necessary to reposition the AFM probe on the probe holder to obtain a maximum sum signal even if step 4.5 is repeated.</li>
</ol>
5. Calibrating the Deflection Sensitivity and Spring Constant of the AFM Probe
<ol>
	<li>Make a scratch on a 60-mm culture dish with a forcep and fill the dish with distilled water. Place the culture dish on the sample holder plate. In this case, mount the dish on the motorized XY scan stage of an inverted microscope.</li>
	<li>Place a magnetic plate holder on top of the plate to keep it secure during imaging.</li>
	<li>Click &#39;Up&#39; arrow in the Navigation menu on the software to move the AFM head up to nearly its highest position to ensure that the AFM probe will not touch the dish. 
	NOTE: This step is very important as it avoids breaking the AFM tip.</li>
	<li>Place the AFM head on the microscope stage.</li>
	<li>Use the microscope to focus on the scratch on the plate. Use the joystick, or click the &#39;Down&#39; arrow in the Navigation menu, to slowly and carefully lower the AFM probe into the PBS. Position the head slightly above the scratch in the plate. 
	NOTE: If the initial position of the AFM head is closer to the sample, the engage time will be significantly reduced.</li>
	<li>Before engaging, re-adjust the position of the laser beam on the AFM tip and the position of the photodiode to obtain the maximum SUM signal as necessary.</li>
	<li>Select a tip type by clicking Change Probe in the Setup menu.</li>
	<li>Click Check Parameters and set the parameters: Scan Size to 0, Scan Rate to 1 Hz, Sample/line to 256 and Deflection Setpoint to 20 nm. Click Engage. The Engage Status window will remain open until the probe makes contact with the surface of the plate. After engaging the probe, click Ramp.</li>
	<li>Click Expanded Mode in the Scan Toolbar menu and set the parameters: Ramp Size to between 500 nm and 1 &#181;m, Ramp Rate to 2 Hz, Number of Samples to 256, Tip Radius to 500 nm, Sample Poisson&#39;s Ratio to 0.5 (assumes the material to be measured is perfectly incompressible), Tip Half Angle to 0, Trigger Mode to Relative and Trigger Threshold to between 20 and 100 nm.</li>
	<li>Click Ramp Continuous in the Ramp Toolbar menu to obtain a force curve on the plate (Figure 3A). In this force curve, set channel 1 to deflection error. 
	NOTE: This will allow the software to graph the results as vertical deflection vs. z position.
	<ol>
		<li>Click at the left or right ends of the force curve and drag the cursor to encompass a linear region of the force curve (Figure 3A, dashed red lines). Use these two lines to mark the boundaries of the sloped region to be fit with a straight line.</li>
	</ol>
	</li>
	<li>Select Ramp in the Toolbar and click Update Sensitivity. Record the value and repeat this step four more times. Select Calibrate in the Toolbar and then click Detector. Input an average value from the five measurements in the Deflection Sensitivity box.</li>
	<li>Click Withdraw 2 - 3 times to raise the AFM head. This step is important to prevent interaction between the AFM tip and the plate during the thermal tune process. Click Thermal Tune in the Toolbar.</li>
	<li>Set Thermal Tune Range of 1 - 100 kHz (This information can be obtained from the manufacturer&#39;s catalog) and Deflection Sensitivity Correction to 1.144 for v-shaped cantilevers and 1.106 for rectangular cantilevers.</li>
	<li>In the Thermal Tune menu, click Acquire Data to obtain a thermal tune curve (Figure 3B). Using the AFM software, place red dashed lines on either side of the curve (as shown in Figure 3B) by dragging the mouse in from the edge of the graph to define the boundaries for fitting. Select either the Lorentzian (Air) or the Simple Harmonic oscillator (Fluid) model depending on which better fits the data.</li>
	<li>Click Fit Data, then Calculate Spring K and save this value. Repeat the spring constant calibrations several times and use the average value. Click Withdraw several times.</li>
	<li>Remove the AFM head from the microscope stage and place it on the alignment unit. Remove the dish from the microscope stage.</li>
</ol>
<img alt="Figure 3" src="/files/ftp_upload/54630/54630fig3.jpg" /> 
Figure 3: AFM Force Curves Used in the Calibration of AFM Probes. (A) A representative AFM force curve (a calibration curve). The extension portion of the force curve between the vertical red dashed lines was used to determine the cantilever deflection sensitivity. (B) A simple harmonic oscillator fit graph used to calculate the spring constant of the cantilever as previously described20. <a href="https://www.jove.com/files/ftp_upload/54630/54630fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Measuring the Elastic Modulus on Mouse Arteries Ex Vivo
<ol>
	<li>Place the 60-mm culture dish containing the aortic tissue on the microscope stage, and secure the plate with the magnetic plate holder.</li>
	<li>Place the AFM head on the microscope stage. Make certain that the AFM probe does not make contact with the aorta but gently makes contact with PBS and forms a meniscus. If necessary, place the AFM head on the alignment unit and click Withdraw until there is enough clearance to prevent contact, then place the AFM head on the microscope stage.</li>
	<li>Click Navigate. Using the joystick or clicking &#39;Down&#39; arrow, slowly lower the AFM probe to locate the cantilever above the aorta. Adjust the photodiode position using the detector positioning knobs on the AFM head if necessary.</li>
	<li>Click Engage to allow the cantilever to make contact with the aorta.</li>
	<li>Once the aorta is engaged, ensure that the piezo center is stable and click Ramp. If the piezo center is fluctuating, this may be the result of a false engagement. Withdraw the probe and increase the vertical offset of the laser on the photodiode by small increments (~ 0.5 V) and re-engage.
	<ol>
		<li>Also, if the probe is far above the sample surface, manually lower the probe closer to the sample surface before re-engaging. If the previous steps do not eliminate the fluctuation, try exchanging the probe.</li>
	</ol>
	</li>
	<li>Set the Ramp Size to 3 &#181;m, the Trigger Mode to Relative and the Trigger Threshold to 100 nm and click Ramp Continuous. Observe that the point of contact between the probe and the sample occurs at approximately the center to the lower &#190; of the Z ramp cycle. If this is not observed, disengage from the sample, adjust the Ramp size as needed, and re-engage the sample.</li>
	<li>Click Microscope in the Menu Bar and then Engage Settings. Change the SPM Withdraw to 30 &#181;m. Click Ramp Continuous in the Ramp Toolbar menu.</li>
	<li>After observing the force curve, click Capture in the Menu bar and then Capture Filename. Enter a desired file name with the ending &#34;.000&#34; (this will allow additional captured files to increase in number by 1). Select a designated-folder and save the data.</li>
	<li>Click Capture in the Capture Toolbar menu. Click Withdraw and then Navigate. Use the Joystick or software controls to position the cantilever over another area of the aorta (next to the initially measured point; see Figure 4).</li>
	<li>Click Engage, Ramp, and Ramp Continuous, respectively. After observing the force curve, click Stop and then Capture.</li>
	<li>Repeat steps 6.9 - 6.10. Capture at least 5 measurements per area as shown in Figure 4. NOTE: We typically collect 15 - 25 force curves from 3 - 5 different locations in each tissue as shown in Figure 4, respectively.</li>
</ol>
<img alt="Figure 4" src="/files/ftp_upload/54630/54630fig4.jpg" /> 
Figure 4: Cartoon of a Cantilever Approaching and Indenting the Tissue (Area 1). This AFM measurement is repeated up to 15 - 25 times from 3 - 5 different locations (Areas 1 - 5) in each artery to acquire the stiffness of the overall tissue sample. <a href="https://www.jove.com/files/ftp_upload/54630/54630fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
7. Data Analysis
<ol>
	<li>Open force curve analysis software.</li>
	<li>Click Find and Open in the menu bar, and double click on a file to be analyzed.</li>
	<li>Click Modify Force Parameters in the toolbar menu to check the deflection sensitivity, Spring Constant, Tip Radius, Tip Half Angle and Poisson&#39;s ratio. To change the parameters check the box next to it and enter the correct values in the new value column. Then click Execute.</li>
	<li>If force curves are noisy, click Boxcar Filter and set inputs for Direction to Extend, Average Points to 3 and Filter Order to 0th. Click Execute to smooth the data.</li>
	<li>Click Baseline Correction in the menu bar and set inputs for Direction to Extend, Plot Units to Force, Type to Separation, Correction Order to 1st and Extend Baseline Source to Extend. Adjust the vertical dotted blue lines to encompass the flat portion of force curve and then click Execute.</li>
	<li>Click Indentation in the toolbar menu and set inputs for Active Curve to Extended, Fit Method to Contact Point Based, Contact Point Algorithm to Treat as Fit Variable, Include Adhesion Force to No, Max Force Fit Boundary to 30%, Min Force Fit Boundary to 0% and Fit Model to Hertzian (Spherical). 
	NOTE: In our measurements, significant adhesion between the tip and sample (a negative deflection of the cantilever in the retraction curve) was rarely observed. In cases where such adhesion is observed, the appropriate model (DMT or JKR) should be used for analysis18,19.</li>
	<li>Save the values of Young&#39;s modulus. Analyze the Young&#39;s (elastic) modulus of each of the 3 - 5 areas (5 force curves per area) taken within a small distance of each other as shown in Figure 4. 
	NOTE: To remove artifacts, exclude Young&#39;s moduli &#62; 100 kPa (normally ~10% of total measurements) from the analysis.</li>
	<li>Calculate a mean Young&#39;s modulus value for each of the 3 - 5 areas (that have been measured) and use these values to calculate a mean elastic modulus for the aorta as a whole. 
	NOTE: At least 3 and more often 4 - 6 individual aortas are used to obtain an accurate assessment of arterial stiffness. The final results are plotted as mean + SEM of &#34;n&#34; independent experiments. The exact number of aortas needed will, of course, depend on the level of accuracy required.</li>
</ol>

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C., K&#228;s, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+Probe-Based+Frequency-Dependent+Microrheology+of+Polymer+Gels+and+Biological+Cells.">Scanning Probe-Based Frequency-Dependent Microrheology of Polymer Gels and Biological Cells.</a> Phys Rev Lett. 85, 880-883 (2000).</li><li>Amin, M., Le, V. P., Wagenseil, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+Testing+of+Mouse+Carotid+Arteries:+from+Newborn+to+Adult.">Mechanical Testing of Mouse Carotid Arteries: from Newborn to Adult.</a> J Vis Exp. , e3733 (2012).</li><li>Laurent, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expert+consensus+document+on+arterial+stiffness:+methodological+issues+and+clinical+applications.">Expert consensus document on arterial stiffness: methodological issues and clinical applications.</a> Eur Heart J. 27, 2588-2605 (2006).</li><li>Plodinec, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+nanomechanical+signature+of+breast+cancer.">The nanomechanical signature of breast cancer.</a> Nat Nano. 7, 757-765 (2012).</li><li>Raman, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+nanomechanical+properties+of+live+cells+using+multi-harmonic+atomic+force+microscopy.">Mapping nanomechanical properties of live cells using multi-harmonic atomic force microscopy.</a> Nat Nano. 6, 809-814 (2011).</li></ol>

The authors have nothing to disclose.

AFM indentation can be used to characterize the stiffness (elastic modulus) of cells and tissues. In this paper, we provide detailed step-by-step protocols to isolate the descending aorta and aortic arch in the mouse and determine the elastic moduli of these arterial regions ex vivo. We now summarize and discuss the technical issues and limitations of the method described in this paper.
Several technical issues can arise in the isolation and analysis of mouse aortas given their small ...

The biomechanical properties of arteries are a critical determinant in cardiovascular disease (CVD) and aging. Arterial stiffness, a major cholesterol independent risk factor and an indicator for the progression of CVD, increases with vascular injury, atherosclerosis, age, and diabetes1-8. Arterial wall stiffening is associated with increased dedifferentiation, migration, and proliferation of vascular smooth muscle cells9-12. In addition, increased arterial stiffness has been linked to enhanced macrophage adhesion1, endothelial permeability and leukocyte transmigration13, and vessel wall remodeling14,15. Thus, therapies that could prevent arterial stiffening in CVD or aging might complement currently available pharmacological interventions that treat CVD by reducing high blood cholesterol. 
AFM is a powerful analytical tool used for various physical and biological applications. AFM is increasingly used to obtain the high-resolution images and characterize the biomechanical properties of soft biological samples such as tissues and cells1,2,10,16,17 with a great degree of accuracy at nanoscale levels. A major advantage of AFM is the fact that it can be used with living cells.
This paper describes our method for measuring the elastic modulus of mouse arteries ex vivo using AFM. The described method shows how we 1) properly isolate mouse arteries (descending aorta and aortic arch) and 2) measure the elastic modulus of these tissues by AFM. Measurements of unloaded elastic moduli in arteries can help to elucidate changes in the extracellular matrix (ECM) that occur in response to vascular injury, CVD, and aging.

<table><tbody><tr><td>BioScope Catalyst AFM system</td><td>Bruker</td><td></td><td></td></tr><tr><td>Nikon Eclipse TE 200 inverted microscope</td><td>Nikon Instruments</td><td></td><td></td></tr><tr><td>Silicon nitride AFM probe</td><td>Novascan Technologies</td><td>PT.SI02.SN.1</td><td>0.06 N/m cantilever; 1 &amp;micro;m SiO&lt;sub&gt;2&lt;/sub&gt; particle</td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>11251-10</td><td>See section 1.4</td></tr><tr><td>Dumont #5SF forceps</td><td>Fine Science Tools</td><td>11252-00</td><td>See section 1.8</td></tr><tr><td>Fine Scissors-ToughCut</td><td>Fine Science Tools</td><td>14058-11</td><td>See section 1.4 (medium sized)</td></tr><tr><td>Vannas-T&amp;uuml;bingen spring scissors</td><td>Fine Science Tools</td><td>15008-08</td><td>See section 1.6 (small sized)</td></tr><tr><td>60 mm TC-treated cell culture dish</td><td>Corning</td><td>353004</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate-Buffered Saline, 1x</td><td>Corning</td><td>21-031-CM</td><td>Without calcium and magnesium</td></tr><tr><td>Krazy Glue instant all purpose liquid</td><td>Krazy Glue</td><td>KG58548R</td><td>See section 2.2</td></tr><tr><td>Gel-loading tips, 1 - 200 &amp;micro;l</td><td>Fisher</td><td>02-707-139</td><td>See section 2.2</td></tr><tr><td>Tip Tweezers</td><td>Electron Microscopy Sciences</td><td>78092-CP</td><td>See section 3.2</td></tr><tr><td>50-mm, clear wall glass bottom dishes</td><td>TED PELLA</td><td>14027-20</td><td>See section 4.4</td></tr></tbody></table>

measuring the stiffness of ex vivo mouse aortas using atomic force microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Figure 5A shows a phase contrast image of the descending (thoracic) aorta from a 6-month old, male C57BL/6 mouse. The AFM cantilever is in place directly above the tissue and ready for indentation. Figures 5B and 5C demonstrate representative force curves obtained by AFM indentation in contact mode. Green lines shown in Figures 5B and 5C represent the best fit curves obtained using the Hertzian model for a sphere. In Figure 5D</s...

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Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

measuring-stiffness-ex-vivo-mouse-aortas-using-atomic-force

Research

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Biology

10.5K Views.  University of Pennsylvania. We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Video: Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Targeted Expression of GFP in the Hair Follicle Using Ex Vivo Viral Transduction

There are many cell types in the hair follicle, including hair matrix cells which form the hair shaft and stem cells which can initiate the hair shaft during early anagen, the growth phase of the hair cycle, as well as pluripotent stem cells that play a role in hair follicle growth but have the potential to differentiate to non-follicle cells such as neurons. These properties of the hair follicle are discussed. The various cell types of the hair follicle are potential targets for gene therapy. Gene delivery system for the hair follicle using viral vectors or liposomes for gene targeting to the various cell types in the hair follicle and the results obtained are also discussed.

The hair follicle is a highly complex appendage of the skin containing a multiplicity of cell types. The follicle undergoes constant cycling through the life of the organism including growth and resorption with growth dependent on specific stem cells. The targeting of the follicle by genes and stem cells to change its properties, in particular, the nature of the hair shaft is, discussed. Hair follicle delivery systems are described, such as liposomes and viral vectors for gene therapy. The nature of the hair follicle stem cells is discussed, in particular, its pluripotency.

There are many cell types in the hair follicle, including hair matrix cells which form the hair shaft and stem cells which can initiate the hair shaft ...

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, ...

Bioengineering

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound and aortic blood pressure is measured invasively with a solid-state pressure catheter. Blood pressure is raised then lowered incrementally by intravenous infusion of vasoactive drugs phenylephrine and sodium nitroprusside. Aortic diameter is measured for each pressure step to characterize the pressure-diameter relationship of the ascending aorta. Stiffness indices derived from the pressure-diameter relationship can be calculated from the data collected. Calculation of arterial compliance is described in this protocol.
This technique can be used to investigate mechanisms underlying increased aortic stiffness associated with cardiovascular disease and aging. The technique produces a physiologically relevant measure of stiffness compared to ex vivo approaches because physiological influences on aortic stiffness are incorporated in the measurement. The primary limitation of this technique is the measurement error introduced from the movement of the aorta during the cardiac cycle. This motion can be compensated by adjusting the location of the probe with the aortic movement as well as making multiple measurements of the aortic pressure-diameter relationship and expanding the experimental group size.

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

We describe a technique for measuring aortic stiffness from its pressure-diameter relationship in vivo in mice. Aortic diameter is recorded by ultrasound and aortic pressure is measured invasively with a solid-state pressure catheter. Blood pressure is changed incrementally and the resulting diameter is measured.

We present a protocol for measuring in vivo aortic stiffness in mice using high-resolution ultrasound imaging. Aortic diameter is measured by ultrasound ...

Medicine

Measuring the Bending Stiffness of Bacterial Cells Using an Optical Trap

We developed a protocol to measure the bending rigidity of filamentous rod-shaped bacteria. Forces are applied with an optical trap, a microscopic three-dimensional spring made of light that is formed when a high-intensity laser beam is focused to a very small spot by a microscope's objective lens. To bend a cell, we first bind live bacteria to a chemically-treated coverslip. As these cells grow, the middle of the cells remains bound to the coverslip but the growing ends are free of this restraint. By inducing filamentous growth with the drug cephalexin, we are able to identify cells in which one end of the cell was stuck to the surface while the other end remained unattached and susceptible to bending forces. A bending force is then applied with an optical trap by binding a polylysine-coated bead to the tip of a growing cell. Both the force and the displacement of the bead are recorded and the bending stiffness of the cell is the slope of this relationship.

We present a protocol for bending filamentous bacterial cells attached to a cover-slip surface with an optical trap to measure the cellular bending stiffness.

We developed a protocol to measure the bending rigidity of filamentous rod-shaped bacteria. Forces are applied with an optical trap, a microscopic ...

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3. On the macro scale the stiffness of tissues and organs within the human body span several orders of magnitude4. Much less is known about how stiffness varies spatially within tissues, and what the scope and spatial scale of stiffness changes are in disease processes that result in tissue remodeling. To better understand how changes in matrix stiffness contribute to cellular physiology in health and disease, measurements of tissue stiffness obtained at a spatial scale relevant to resident cells are needed. This is particularly true for the lung, a highly compliant and elastic tissue in which matrix remodeling is a prominent feature in diseases such as asthma, emphysema, hypertension and fibrosis. To characterize the local mechanical environment of lung parenchyma at a spatial scale relevant to resident cells, we have developed methods to directly measure the local elastic properties of fresh murine lung tissue using atomic force microscopy (AFM) microindentation. With appropriate choice of AFM indentor, cantilever, and indentation depth, these methods allow measurements of local tissue shear modulus in parallel with phase contrast and fluorescence imaging of the region of interest. Systematic sampling of tissue strips provides maps of tissue mechanical properties that reveal local spatial variations in shear modulus. Correlations between mechanical properties and underlying anatomical and pathological features illustrate how stiffness varies with matrix deposition in fibrosis. These methods can be extended to other soft tissues and disease processes to reveal how local tissue mechanical properties vary across space and disease progression.

The stiffness of the extracellular matrix strongly influences multiple behaviors of adherent cells. Matrix stiffness varies spatially throughout a tissue, and undergoes modification in various disease conditions. Here we develop methods to characterize spatial variations in stiffness in normal and fibrotic mouse lung tissue using atomic force microscopy microindentation.

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3.  On the macro scale the stiffness of tissues and organs ...

Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 times of its original size. Clinical and experimental studies have shown that small aneurysms may rupture, while a subpopulation of large aneurysms may remain stable. Thus, in addition to the measurement of intraluminal diameter of the aorta, knowledge of structural traits of the vessel wall may provide important information to assess the stability of the AAA. Aortic stiffening has recently emerged as a reliable tool to determine early changes in the vascular wall. Pulse propagation velocity (PPV) along with the distensibility and radial strain are highly useful ultrasound-based methods relevant for assessing aortic stiffness. The primary purpose of this protocol is to provide a comprehensive technique for the use of ultrasound imaging system to acquire images and analyze the structural and functional properties of the aorta as determined by MILD, PPV, distensibility and radial strain.

This manuscript describes a detailed protocol for using high frequency ultrasound imaging to measure luminal diameter, pulse propagation velocity, distensibility and radial strain on a mouse model of abdominal aortic aneurysm.

An abdominal aortic aneurysm (AAA) is defined as a localized dilation of the abdominal aorta that exceeds the maximal intraluminal diameter (MILD) by 1.5 ...

Immunology and Infection

Measurement of Endothelium-Dependent Vasorelaxation in the Mouse Thoracic Aorta Using Tensometric Small Volume Chamber Myography

Small volume chamber tensometric myography is a commonly used technique to evaluate the vascular contractility of small and large blood vessels in laboratory animals and small arteries isolated from human tissue. The technique allows researchers to maintain isolated blood vessels in a tightly controlled and standardized (near-physiological) setting, with the option of adjusting to various environmental factors, while challenging the isolated vessels with different pharmacological agents that can induce vasoconstriction or vasodilation. The myograph chamber also provides a platform to measure vascular reactivity in response to various hormones, inhibitors, and agonists that may impact the function of smooth muscle and endothelial layers separately or simultaneously. The blood vessel wall is a complex structure consisting of three different layers: the intima (endothelial layer), media (smooth muscle and elastin fibers), and adventitia (collagen and other connective tissue). To gain a clear understanding of the functional properties of each layer, it is critical to have access to an experimental platform and system that would allow for a combinational approach to study all three layers simultaneously. Such an approach demands access to a semi-physiological condition that would mimic the in vivo environment in an ex vivo setting. Small volume chamber tensometric myography has provided an ideal environment to evaluate the impact of environmental cues, experimental variables, or pharmacological agonists and antagonists on vascular properties. For many years, scientists have used the tensometric myograph technique to measure endothelial function and smooth muscle contractility in response to different agents. In this report, a small volume chamber tensometric myograph system is used to measure endothelial function in the isolated mouse aorta. This report focuses on how small volume chamber tensometric myography can be used to evaluate the functional integrity of the endothelium in small segments of a large artery such as the thoracic aorta.

The present protocol describes the concepts and technical application of the tensometric myograph technique using a multi-chamber myograph system in the experimental ex vivo assessment of mouse aortic endothelial function.

Small volume chamber tensometric myography is a commonly used technique to evaluate the vascular contractility of small and large blood vessels in ...

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell behavior such as cell function, differentiation, and motility. However, as these processes are not homogeneous throughout the whole organ, it has become increasingly important to understand changes in the mechanical properties of tissues on the cellular level.
To be able to monitor the stiffening of collagen-rich areas within the liver lobes, this paper presents a protocol for measuring liver tissue elastic moduli with high spatial precision by atomic force microscopy (AFM). AFM is a sensitive method with the potential to characterize local mechanical properties, calculated as Young&#39;s (also referred to as elastic) modulus. AFM coupled with polarization microscopy can be used to specifically locate the areas of fibrosis development based on the birefringence of collagen fibers in tissues. Using the presented protocol, we characterized the stiffness of collagen-rich areas from fibrotic mouse livers and corresponding areas in the livers of control mice.
A prominent increase in the stiffness of collagen-positive areas was observed with fibrosis development. The presented protocol allows for a highly reproducible method of AFM measurement, due to the use of mildly fixed liver tissue, that can be used to further the understanding of disease-initiated changes in local tissue mechanical properties and their effect on the fate of neighboring cells.

We present a protocol to measure the elastic moduli of collagen-rich areas in normal and diseased liver using atomic force microscopy. The simultaneous use of polarization microscopy provides high spatial precision for localizing collagen-rich areas in the liver sections.

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell ...

Measurement of Stiffness in Mouse Retinal Capillaries and Subendothelial Matrix

Isolation of Mouse Retinal Capillaries and Subendothelial Matrix for Stiffness Measurement Using Atomic Force Microscopy

Retinal capillary degeneration is a clinical hallmark of the early stages of diabetic retinopathy (DR). Our recent studies have revealed that diabetes-induced retinal capillary stiffening plays a crucial and previously unrecognized causal role in inflammation-mediated degeneration of retinal capillaries. The increase in retinal capillary stiffness results from the overexpression of lysyl oxidase, an enzyme that crosslinks and stiffens the subendothelial matrix. Since tackling DR at the early stage is expected to prevent or slow down DR progression and associated vision loss, subendothelial matrix, and capillary stiffness represent relevant and novel therapeutic targets for early DR management. Further, direct measurement of retinal capillary stiffness can serve as a crucial preclinical validation step for the development of new imaging techniques for non-invasive assessment of retinal capillary stiffness in animal and human subjects. With this view in mind, we here provide a detailed protocol for the isolation and stiffness measurement of mouse retinal capillaries and subendothelial matrix using atomic force microscopy.

Author Spotlight: Investigating Vascular Stiffness and Inflammation in Early Diabetic Retinopathy

We recently identified retinal capillary stiffening as a new paradigm for retinal dysfunction associated with diabetes. This protocol elaborates the steps for isolation of mouse retinal capillaries and the subendothelial matrix from retinal endothelial cultures, followed by a description of the stiffness measurement technique using atomic force microscopy.

Retinal capillary degeneration is a clinical hallmark of the early stages of diabetic retinopathy (DR). Our recent studies have revealed that ...

Mechanical Characterization of Tissue Stiffness During Axolotl Limb Regeneration

Atomic Force Microscopy Measurements of Cartilage in Intact and Regenerating Axolotl Limbs

Mechanical forces provide important signals for normal cell function and pattern formation in developing tissues, and their role has been widely studied during embryogenesis and pathogenesis. Comparatively, little is known of these signals during animal regeneration. 
The axolotl is an important model organism for the study of regeneration, given its ability to fully restore many organs and tissues after injury, including missing cartilage and bone. Due to its crucial role as the main supporting tissue in the vertebrate body, regaining skeletal function during regeneration requires both the restoration of the missing structures as well as their mechanical properties. This protocol describes a method for processing axolotl limb samples for atomic force microscopy (AFM), which is the gold standard for probing cell and tissue mechanical properties at high spatial resolution. 
Taking advantage of the regenerative capabilities of the axolotl, this study measured the stiffness of limb cartilage during homeostasis and two stages of limb regeneration: tissue histolysis and cartilage condensation. We show that AFM is a valuable tool for gaining insights into dynamic tissue restructuring and the mechanical changes that occur during regeneration.

Author Spotlight: Exploring the Role of Mechanical Signals in Tissue Regeneration Through Atomic Force Microscopy 

In this protocol, we show how to prepare axolotl tissue for atomic force microscopy (AFM) and perform indentation measurements in intact and regenerating limb cartilage.

Mechanical forces provide important signals for normal cell function and pattern formation in developing tissues, and their role has been widely studied ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

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Targeted Expression of GFP in the Hair Follicle Using Ex Vivo Viral Transduction

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Measuring Ascending Aortic Stiffness In Vivo in Mice Using Ultrasound

Measuring the Bending Stiffness of Bacterial Cells Using an Optical Trap

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Measurement of Pulse Propagation Velocity, Distensibility and Strain in an Abdominal Aortic Aneurysm Mouse Model

Measurement of Endothelium-Dependent Vasorelaxation in the Mouse Thoracic Aorta Using Tensometric Small Volume Chamber Myography

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy

Isolation of Mouse Retinal Capillaries and Subendothelial Matrix for Stiffness Measurement Using Atomic Force Microscopy

Atomic Force Microscopy Measurements of Cartilage in Intact and Regenerating Axolotl Limbs