We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus 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.
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.
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
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. Please click here to view a larger version of this figure.
2. Preparing Tissue Samples for the AFM Measurements
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. Please click here to view a larger version of this figure.
3. Loading the Probe
4. Aligning the Laser on the Probe
5. Calibrating the Deflection Sensitivity and Spring Constant of the AFM Probe
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. Please click here to view a larger version of this figure.
6. Measuring the Elastic Modulus on Mouse Arteries Ex Vivo
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. Please click here to view a larger version of this figure.
7. Data Analysis
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, the mean stiffness of the descending aortas and aortic arches were determined from three individual mice as described in the text. Note that the descending aorta and aortic arch represent regions of laminar and disturbed flow, respectively. Nevertheless, their (mechanically unloaded) elastic moduli are similar as determined by AFM.
Figure 5: The Analyzed Force-Indentation Curves and Stiffness Data. (A) Phase contrast micrograph showing an AFM cantilever over a mouse descending aorta ex vivo. Scale bar, 100 µm. (B-C) A set of two representative force-indentation curves acquired for (B) the descending aorta and (C) the aortic arch. (D) Mean stiffness of the mouse descending aorta and aortic arch was determined as described above. Error bars show mean + SEM; n= 3 independent biological replicates. Please click here to view a larger version of this figure.
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 and thin nature. When the cleaned arteries are being opened longitudinally, care must be taken not to disrupt the intima (luminal surface) where the elastic modulus data is collected. Excess PBS in the final sample dish prevents proper gluing of the arteries onto the dish, and this needs to be carefully removed using a paper tissue without touching the artery before gluing. Note, however, that the isolated artery can stick to the paper tissue while attempting to remove excessive PBS, and this can damage the sample. While gluing the arteries to the culture dish, it is extremely important that the artery is not stretched and that only a small amount of adhesive is applied bit-by-bit to each end of the artery. When performing the AFM-indentation, it is critical to avoid probing at the ends of the artery where the adhesive has been applied. Lastly, because freshly isolated arteries are used for the measurements of elastic modulus, the time elapsed from harvesting to measurement should be minimized.
Since the spherical tips used in this study have a radius of 500 nm, an indentation depth of up to 500 nm can be accurately analyzed using the Hertz model. This indentation depth would correlate with the top layer of endothelial cells located in the intima21. To probe the mechanics of the sub-endothelial layers of arterial samples, larger colloidal probes (10 – 20 µm in diameter) could be used for indentation and changes in the calculated stiffness with indentation depth could be determined22,23. Also, since biological samples are usually viscoelastic rather than purely elastic, stress relaxation measurements (monitoring the decay in applied force at a constant indentation depth) could be used to determine the viscous component of arterial mechanical properties24,25.
A potential limitation of AFM analysis is the scan size, which only measures a small region of the tissue. Most biological samples, including arteries and cells, are not topographically and biomechanically homogeneous, and this heterogeneity can potentially result in artifactually large variations of the elastic modulus depending on smoothness of the indentation site, indentation depth, and the amount of force applied to the sample surface. To obtain a representative mean value of the overall elastic modulus, it is important to perform repeated AFM-indentation at multiple random locations through the arteries, as shown in Figure 4. Furthermore, we use an appropriately-sized spherical tip, as previous studies have demonstrated that AFM-indentation with a sharp pyramidal tip resulted in higher estimates of the elastic modulus and damaged soft biological samples26,27.
Even though our ex vivo AFM measurements of arterial stiffness appear to agree well with molecular markers of smooth muscle cell stiffening2, it is important to realize that these arterial samples are not mechanically loaded, and mechanical load is likely to play an important role in overall arterial mechanics. Comparisons of arterial stiffness as determined by AFM, myography (an ex vivo method compatible with mechanical loading)28 and pulse-wave velocity (an in vivo measurement of arterial stiffness)29 will likely provide the most comprehensive understanding of arterial mechanics.
We have described the AFM in contact mode to measure the ex vivo stiffness of mouse arteries. This method estimates the average tissue stiffness by randomly indenting on multiple regions of the aorta. To examine the spatial distribution of stiffness from bigger regions of tissues, a recent paper has used AFM in force-volume mode30. This approach simultaneously obtained a map of stiffness variations and a high-resolution image of topographical surface of the tissue. In addition to measuring the stiffness of tissue samples, the AFM in force-volume mode can be used in cells to monitor the spatial distributions of intracellular stiffness10,31.
The authors have nothing to disclose.
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.
BioScope Catalyst AFM system | Bruker | ||
Nikon Eclipse TE 200 inverted microscope | Nikon Instruments | ||
Silicon nitride AFM probe | Novascan Technologies | PT.SI02.SN.1 | 0.06 N/m cantilever; 1 µm SiO2 particle |
Dumont #5 forceps | Fine Science Tools | 11251-10 | See section 1.4 |
Dumont #5SF forceps | Fine Science Tools | 11252-00 | See section 1.8 |
Fine Scissors-ToughCut | Fine Science Tools | 14058-11 | See section 1.4 (medium sized) |
Vannas-Tübingen spring scissors | Fine Science Tools | 15008-08 | See section 1.6 (small sized) |
60mmTC-treated cell culture dish | Corning | 353004 | |
Dulbecco's Phosphate-Buffered Saline, 1X | Corning | 21-031-CM | Without calcium and magnesium |
Krazy Glue instant all purpose liquid | Krazy Glue | KG58548R | See section 2.2 |
Gel-loading tips, 1-200 µL | Fisher | 02-707-139 | See section 2.2 |
Tip Tweezers | Electron Microscopy Sciences | 78092-CP | See section 3.2 |
50-mm, clear wall glass bottom dishes | TED PELLA | 14027-20 | See section 4.4 |