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Daniel Chaparro
1
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Valentina Dargam
1
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Paulina Alvarez
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Jay Yeung
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Ilyas Saytashev
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Jenniffer Bustillo
Department of Mechanical and Materials
Engineering,
Florida International University,
Miami, FL 33174
Archana Loganathan
Department of Mechanical and Materials
Engineering,
Florida International University,
Miami, FL 33174
Jessica Ramella-Roman
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174
Arvind Agarwal
Department of Mechanical and Materials
Engineering,
Florida International University,
Miami, FL 33174
Joshua D. Hutcheson
2
Department of Biomedical Engineering,
Florida International University,
Miami, FL 33174;
Biomolecular Sciences Institute,
Florida International University,
Miami, FL 33199
e-mail: jhutches@fiu.edu
A Method to Quantify Tensile
Biaxial Properties of Mouse
Aortic Valve Leaflets
Understanding aortic valve (AV) mechanics is crucial in elucidating both the mechanisms
that drive the manifestation of valvular diseases as well as the development of treatment
modalities that target these processes. Genetically modified mouse models have become
the gold standard in assessing biological mechanistic influences of AV development and
disease. However, very little is known about mouse aortic valve leaflet (MAVL) tensile
properties due to their microscopic size (500 lm long and 45 lm thick) and the lack of
proper mechanical testing modalities to assess uniaxial and biaxial tensile properties of
the tissue. We developed a method in which the biaxial tensile properties of MAVL tissues
can be assessed by adhering the tissues to a silicone rubber membrane utilizing dopamine
as an adhesive. Applying equiaxial tensile loads on the tissue–membrane composite and
tracking the engineering strains on the surface of the tissue resulted in the characteristic
orthotropic response of AV tissues seen in human and porcine tissues. Our data suggest
that the circumferential direction is stiffer than the radial direction (n ¼6, P ¼0.0006) in
MAVL tissues. This method can be implemented in future studies involving longitudinal
mechanical stimulation of genetically modified MAVL tissues bridging the gap between
cellular biological mechanisms and valve mechanics in popular mouse models of valve
disease. [DOI: 10.1115/1.4046921]
Keywords: mouse aortic valve mechanics, aortic valve, anisotropy
Introduction
Understanding aortic valve (AV) mechanics is crucial in eluci-
dating both the mechanisms that drive the manifestation of valvu-
lar diseases as well as the development of treatment modalities
that target these processes. Orthogonally aligned elastin and
1
These authors contributed equally.
2
Corresponding author.
Manuscript received January 2, 2020; final manuscript received April 9, 2020 ;
published online August 31, 2020. Assoc. Editor: Anna Grosberg.
Journal of Biomechanical Engineering OCTOBER 2020, Vol. 142 / 100801-1Copyright V
C2020 by ASME
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collagen fibers dictate AV structure and function [1]. Radially
aligned elastin fibers allow the AV leaflets to stretch toward the
center of the AV orifice during diastole, preventing retrograde
blood flow. Circumferentially aligned collagen fibers provide the
necessary tensile strength to prevent leaflet prolapse under this
diastolic load. Alterations in fiber structure and composition
within the AV compromise valve mechanics and thus function
[2]. AV leaflet mechanics have traditionally been quantified using
uniaxial or biaxial tensile testing modalities in which the tensile
stiffness of these tissues in the radial and circumferential direc-
tions can be determined [3–5]. Studies including these tests have
extensively reported the characteristic orthotropic response of AV
leaflets in which the radial direction is softer than the circumferen-
tial direction.
Genetically modified mouse models recapitulating certain
aspects of human AVs have facilitated mechanistic studies of AV
development, congenital defects, and disease progression [6].
However, as convenient as these mouse models are in clarifying
some of the biological mechanisms behind AV development and
disease progression, very little is known about the gross mechani-
cal properties of the mouse AV due to its microscopic size (500
lm long and 45 lm thick) [7]. As such, traditional uniaxial and
biaxial tensile testers cannot be implemented to mouse aortic
valve leaflet (MAVL) tissues.
Biomechanical analyses of MAVLs have traditionally
employed local strain measurement techniques such as atomic
force microscopy, nano-indentation, and micropipette aspiration
[8–10]. While these techniques may be used to quantify regional
differences in MAVL tissue stiffness, they displace MAVL tissues
perpendicular to the radial/circumferential plane. Therefore, these
techniques do not recreate the physiologically relevant loads and
displacements experienced by AV leaflets in-vivo and are also
unable to determine anisotropic tensile properties of the tissue.
Physiologically relevant mechanical testing of MAVL tissues that
recapitulate biaxial tests performed in human and porcine valves
are needed to better understand the relationship between human
and mouse AVs and connect the biomolecular insight provided by
mouse models to AV biomechanics [11].
In this study, we developed a method to determine MAVL ani-
sotropic tensile properties by adhering the tissue to a silicone
membrane coated with dopamine. A Flexcell system was used to
induce equiaxial tensile loads on the tissue–membrane composite,
and a custom MATLAB script was used to determine the engineering
strains in the radial and circumferential directions on the leaflet
surface. We demonstrate anisotropic MAVL behavior that mimics
previous observations from human and porcine AV tensile testing.
Methodology
Dopamine Coating and Tissue Preparation. StageFlexer
(Flexcell International, Burlington, NC) membranes were treated
with dopamine (A11136 dopamine hydrochloride 99%, Alfa
Fig. 1 Preparation of MAVL on dopamine-coated membrane: (a) adult WT (C57/BL6) mouse heart,
(b) intact aortic root postremoval of surrounding tissues, and (c) the valve is cut open at L–N coro-
nary commissure to maintain leaflet integrity, thus exposing the leaflets. Cusps are resected by
cutting close to the base of the leaflet. (d) The leaflet is then placed flat on a dopamine-coated
StageFlexer
V
R
silicone rubber membrane.
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Aesar, Haverhill, MA) (0.1, 0.3, 0.5 mg/mL) in 0.01 M tris buffer
solution and left at room temperature overnight (18h). Polydop-
amine has been shown to promote adhesion of biological samples
to synthetic surfaces [12,13]. Polydopamine coated surfaces have
shown to reduce substrate surface hydrophobicity and promote in-
vitro tissue development [14]. Therefore, dopamine solutions were
used to promote adhesion between substrate surface and tissue.
C57/BL6 wild type (WT) mouse hearts were transported to the
laboratory in ice cold 1phosphate buffered saline (PBS) where
a stereoscopic zoom microscope (Nikon SMZ645) was used to
dissect the hearts and resect the AV leaflets (Fig. 1). Tissues were
periodically rinsed with ice cold 1PBS to prevent drying. A
transverse cut was made to remove the apex of the heart (Fig.
1(a)) and expose the left and right ventricles, then the atria and
pulmonary artery were carefully removed proximal to the aortic
root (Fig. 1(b)). The AV was then opened by cutting the commis-
sure between the left and noncoronary (L–N) leaflets from the
ventricular side and through the remaining mitral valve leaflet tis-
sue (Fig. 1(c)). Each MAVL was resected by cutting at the aortic
root base. MAVL tissues were carefully flattened onto the
dopamine-coated membrane surface by placing a leaflet in a
3lL1PBS droplet on the membrane and carefully aspirating
the solution while simultaneously manipulating the leaflet until
proper contact throughout the tissue surface was obtained (Fig.
1(d)).
Tissue Adhesion Verification. The Flexcell StageFlexer sys-
tem could not be used to measure tissue–membrane adhesion
since the maximum allowable strain (15%) did not induce tissue
detachment. Therefore, tissue–membrane adhesion was tested by
conducting uniaxial strain tests with an MTI SEMtester (MTI
instruments, Albany, NY) on the composite to determine the strain
at which noticeable separation occurred. To assess the effect of
dopamine concentration on tissue adhesion, two dopamine con-
centrations were tested, 0.1 and 0.5 mg/mL (n¼1). After proper
tissue adhesion, the surface of the membrane containing the
MAVL is sprayed with red tissue marking dye while the other
side is sprayed with blue acrylic paint. This aids in determining
the time of interest (TOI), defined here as the point throughout the
tensile test where noticeable detachment occurs evident by a dif-
ference in strain between the red and blue fiducial markers. Rec-
tangular strips (15 7mm
2
) of the tissue-containing-membrane
are cut out such that the leaflet resides near the center of the rec-
tangular strip with the radial direction aligned parallel to the pri-
mary axis of strain (Fig. 2(a)). Uni-axial tension was applied and
quantified using the MTI SEMtester (100 Hz sampling rate) while
simultaneously recording the leaflet with a microscope video cam-
era (30 frames per second) to determine the TOI. TOI was man-
ually determined by scrubbing through video footage of the
experiment and noting the grip to grip strain level at these time
points.
Fig. 2 Experimental setup for uniaxial and equiaxial testing of tissue–membrane composite: (a)
tissue–membrane adhesion verification was tested by applying uniaxial tension to the sample,
consisting of a rectangular strip of the membrane with the adhered leaflet, using an MTI SEMtester
at room temperature with constant 13PBS washes to prevent the tissue from drying, (b) Stage-
Flexer dopamine coated membrane with sample, depicting the location of the leaflet on the loading
station, and (c) the leaflet circumferential and radial orientations were manually determined
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Mouse Aortic Valve Leaflet Strain Testing. Mouse aortic
valve leaflets were adhered near the center of the silicon rubber
membrane coated with a 0.3 mg/mL dopamine solution ensuring
proper adhesion and minimizing black dopamine precipitants at
the higher 0.5 mg/mL concentration (n¼6). Dopamine is an
organic chemical that interacts with slightly basic pH solutions by
a spontaneous oxidative reaction to form a layer of polydopamine,
a type of melanin [14]. The silicone membrane was then loaded
into a StageFlexer Strain Device and 1PBS was added to pre-
vent drying (Fig. 2(b)). When a vacuum is applied, an equiaxial
load is applied as the membrane deforms over a lubricated loading
post [15]. Ten preconditioning cycles of 52.86 kPa load at 0.5 Hz
were conducted prior to image acquisition. Microscopic images of
the leaflet were acquired at five different vacuum pressures loads
(6.72, 15.62, 26.44, 38.93, and 52.86 kPa) using a FlexCell system
(Flexcell FX-5000), resulting in five load–strain configurations.
Each vacuum load was applied for 30 s, allowing for reposition-
ing, refocusing, and preventing strain rate-dependent effects.
One image per load configuration of each MAVL sample was
acquired using an upright microscope (Zeiss Axioscope Upright
Fluorescent Microscope, Oberkochen, Germany) with a mono-
chromatic Axiocam attachment. IMAGEJ was used in conjunction
with a “bio-formats” plug-in to display the microscope image data
files. In the context of this report, radial and circumferential direc-
tions refer to the anatomy of aortic valve leaflets. These in-vivo
directions have unique mechanical properties that manifest as ani-
sotropic leaflet behavior measured in biaxial testing. Therefore, we
use the radial and circumferential terminology on the resected leaf-
lets to test this characteristic biaxial behavior as done previously
for aortic valve leaflets from larger mammals. The circumferential
leaflet direction within the acquired image coordinate system is
manually determined and the radial direction is defaulted to be
orthogonal to the circumferential direction (Fig. 2(c)). MAVL tis-
sues have distinct melanocytic pigment patterns, which serve as
trackable markers. Position of multiple markers at each configura-
tion are recorded by using the “multipoint” tool offered by IMAGEJ,
which stores xand ycoordinates of selected markers on the image.
These markers are manually placed on distinct pigment patterns
for consistency throughout configurations. A custom MATLAB script
creates lines and calculates the angles and Euclidean distance
between the permutations of all the markers at each configuration.
Then, strain of each line at its corresponding load configuration is
calculated as ðDL=L0Þ, the engineering strain, with the 6.72 kPa
load configuration serving as the original configuration (L0)to
ensure that the MAVL was under basal tension (i.e., not folded).
Manual selection of the circumferential direction, with respect to
the MAVL anatomical geometry (Fig. 2(c)), is used to determine
the strains that fall within 10 deg of the primary axes of the valve
(radial and circumferential). Stiffness of the tissue–membrane
composite is determined as the slope of a linear fit of the overall
tissue strain–load curves.
Membrane Strain Verification. Verification of isotropic
StageFlexer membrane displacement under the same load regimen
as the tissue–membrane composite was conducted by following
the procedures outlined in the Tissue Strain Testing section with a
single key difference. Tissue marking dye was sprayed on top of
the membrane to serve as trackable fiducial markers on the other-
wise transparent membrane surface.
Tissue Integrity and Fiber Alignment. To assess tissue viabil-
ity, an assay for live/dead cell staining (LIVE/DEAD
TM
viability/
cytotoxicity kit. ThermoFisher Scientific, Waltham, MA) was per-
formed after MAVL tissue mechanical testing. Images were
obtained using the Zeiss fluorescent microscope (n¼3). Images
of calcein-acetoxymethyl and ethidium homodimer fluorescence
can be seen in (Fig. 3(a), green and red, respectively). To verify
MAVL fiber structure after mechanical testing and confirm radi-
ally and circumferentially oriented elastin and collagen fibers,
respectively, two-photon emission fluorescence (TPEF) of elastin
(red) and collagen (cyan) second harmonic generation (SHG)
were obtained (Fig. 3(b)). A home-built laser scanning nonlinear
microscope with a broadband femtosecond excitation laser (Ele-
ment 600, Femtolasers, Austria) was used to obtain SHG images
of collagen and TPEF of elastin. Images were obtained by scan-
ning a laser beam with a pair of galvanometer mirrors (Thorlabs,
Newton, NJ) and directing it into a 20/1 NA water immersion
objective (Olympus, Japan), via a dichroic mirror (655spxr,
Chroma, Bellows Falls, VT) to separate two-photon signals from
a reflected fundamental wavelength. A pair of photomultiplying
tubes (Hamamatsu, Japan) with bandpass filters (400 nm central
wavelength/40 nm bandwidth and 480 nm central wavelength/
40 nm bandwidth) recorded SHG and TPEF signals, respectively.
Over 1000 1000-pixel images were acquired at 0.5 frames per
second, 10 images were averaged to improve the signal to noise
ratio. In postprocessing, a smoothing filter was used with IMAGEJ
(ImageJ, NIH, Bethesda, MD, 2-pixel mean radius filter). Images
were square rooted to improve visibility.
Results
Tissue Adhesion Verification. Mouse aortic valve leaflet tis-
sues on the StageFlexer membranes remained attached throughout
Fig. 3 MAVL tissue integrity and fiber alignment after mechanical stimulation: (a) calcein-
acetoxymethyl (green) fluorescence and ethidium homodimer (red) fluorescence were used to
image live and dead MAVL cells, respectively, (b) two-photon microscopic images of circumferen-
tially, and (c)-aligned collagen (SHG, cyan) and radially(r)-aligned elastin (TPEF, red) fibers
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strain levels far beyond those capable by the FlexCell system
(14.8%). The MTI SEMtester can reach strains of up to 200% with
a starting axial length of 10 mm between the sample grips. Tissue
placed on the 0.5 mg/mL dopamine-coated membrane never
showed signs of detachment while the tissue on the 0.1 mg/mL
dopamine-coated membrane showed signs of detachment at 172%
strain (n¼1) (Table 1).
Membrane Strain Verification. Directional strain was deter-
mined by binning measured lines between markers into 10-deg
angle bins with respect to an arbitrary horizontal (Fig. 4). Similar
to previous observations [15], an angle independence of the strain
field on the isotropic lone silicone membrane indicate an equiaxial
load field (Fig. 4(a)). However, an angle dependence is observed
in the tissue–membrane composite, indicating anisotropic
behavior (Fig. 4(b)).
Mouse Aortic Valve Leaflet Strain. Strain–load curves depict
the mechanical properties of the tissue–membrane composite
(Fig. 5)(n¼6, biological replicates). Equiaxial tensile load on the
tissue–membrane composite resulted in an orthotropic strain
response (Fig. 6). Lone membrane radial stiffness using the same
methodology mentioned earlier in this report was calculated to be
615 kPa (n¼1). The tissue–membrane composite stiffness, calcu-
lated as the mean slope 6standard deviation of a linear fit for
each curve, was 7356162 kPa in the radial direction and
8906153 kPa in the circumferential direction (Figs. 6(a)–6(c)). A
paired T-test confirms a statistically significant difference
(P¼0.0006) between the radial and circumferential strains of
these tissues (Fig. 6(d)). The mean difference between these
orthogonal directions was 154.768.8 kPa.
It is important to note that the term “stiffness” in the develop-
ment of this method is used to describe the resistance to deforma-
tion by the tissue–membrane composite to the applied equiaxial
load. This load is created by pulling a vacuum pressure within the
StageFlexer chamber and is reported in kPa. Since engineering
strain is a dimensionless measure of displacement, the slope of the
linear fit to the load–displacement curves are in the units of kPa as
well. This is not to be confused with the classical measure of stiff-
ness from a standard stress–strain curve. Further development of
this method would include a mathematical model to determine the
Table 1 Tissue adhesion verification. Uniaxial tensile tests
were performed using an MTI SEMtester to identify adhesive
properties of MAVL tissue to silicone rubber membrane using
different dopamine concentrations. The strain at the time of
detachment for both dopamine concentrations is greater than
the applied strain in the experimental equiaxial mechanical test-
ing regimen (10%).
Dopamine concentration (mg/mL) Strain at detachment (%)
0.1 171.6
0.5 >200 (no evidence of detachment)
Fig. 4 Mean-measured strain experienced by the lone membrane and tissue–membrane composite. To measure strain,
markers were manually selected and tracked on the membrane alone and tissue surface of the tissue–membrane composite
for different applied vacuum pressure loads. The Euclidian distance between each marker was used to calculate the engineer-
ing strain. Strain was averaged and binned in 10 deg increments based on the arbitrary horizontal of the image. (a) Tissue
marking dye speckle pattern markers on a silicone rubber membrane were used to verify the strain experienced by the iso-
tropic membrane at each load configuration, indicating an equiaxial load field. (b) Pigment on a MAVL was used as trackable
markers to measure the strain experienced by the tissue–membrane composite at each equiaxial load configuration. An
angle-dependent, anisotropic, behavior is observed.
Fig. 5 Strain–load curve for each MAVL tissue–membrane
composite sample (dash line) and the mean of all samples
(n56, solid line). The stiffness for all samples, as defined by
the slope of the mean strain–load curve, is 7966157 kPa.
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applied stress and decouple the tissue characteristics from the
tissue–membrane composite.
Discussion
This study developed a method in which MAVL biaxial tensile
mechanics can be assessed by applying equiaxial tensile loads on
a tissue–membrane composite while tracking the engineering
strains on the surface of the tissue. Tensile biaxial mechanical
properties of MAVL tissues have not been previously reported;
thus, this method may allow future studies to combine murine-
based mechanistic studies of AV development and disease to
MAVL mechanics. Similar to the characteristic orthotropic
mechanical properties of human and porcine AVs, statistically
significant orthotropic tissue level mechanics depict a stiffer cir-
cumferential direction compared to the radial direction. During
lower load configurations, the circumferential strains are higher
than radial strains, whereas the radial strain becomes higher than
the circumferential strain at higher loads, implicating possible col-
lagen uncrimping of the tissues at intermediate strain levels.
Previous studies have reported a similar response in porcine aortic
valves, where crimped collagen fibers provide little resistance
when small loads are applied but resist deformation once suffi-
cient loads promote uncrimping of the collagen fibers [1].
The strain range provided by the FlexCell system (0–15%) lim-
its the assessment of MAVL mechanics at high strains utilizing
this method. It is possible that the biaxial response is more pro-
found at higher levels of strain as seen with larger mammal mod-
els. As with all soft tissues, MAVL tissues are expected to exhibit
viscoelastic material properties. The nonlinear displacement
response to increasing load may be evident at higher strain values
(>10%) as seen in larger mammal models testing the same tissues
[16]. At the lower strains used in this study (2–8%), the MAVL
tissues exhibit a linear behavior. Further experimentation to deter-
mine the viscoelastic properties of MAVL tissues would necessi-
tate more capable biaxial mechanical testing setups as well as
decoupling the leaflet behavior from the tissue–membrane com-
posite. In future studies, traditional biaxial tensile testers could be
implemented using the tissue–membrane composite to assess the
tissue response at higher levels of strain. Longer timescale
Fig. 6 Strain–load curve for each MAVL tissue–membrane composite sample (dash line) (n56) and the mean of all samples
(solid line), from orthogonal directions on the leaflet surface: (a) circumferential strain–load curve, (b) radial strain–load
curve, (c) the tissue–membrane composite mean stiffness for all samples, as defined by the slope of the mean strain–load
curve, is 7356162 kPa in the radial direction (circles) and 8906153 kPa in the circumferential direction (crosses), and (d) com-
parison of MAVL stiffness in the radial and circumferential strain directions are significantly different (paired t-test, two-tailed,
95% confidence interval, p<0.05)
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experimentation using this method to controllably mechanically
stimulate MAVL tissues would elucidate on the role of mechani-
cal load on the various established mouse models of aortic valve
disease.
Experiments were originally conducted with eight leaflet sam-
ples; however, our analyses indicated that two of the samples did
not remain attached throughout the duration of the mechanical
testing regimen. The two poorly adhered samples were evident by
either (1) visible tissue sections floating in solution (e.g., only the
free edge would attach and the base would float), or (2) tension
was observed in one direction and compression in another (Pois-
son’s effect). If the materials are perfectly bonded, all directions
would be in tension during the testing. The load–strain curves
found by tracking surface markers are only applicable in meas-
uring properties of the tissue–membrane composite if the tissue
and membrane displace equally as a perfectly bonded composite
material. Therefore, the poorly adhered leaflets were excluded from
the study. For the six samples used in this study, perfect adhesion
between the tissue and the membrane was assumed in the analyses.
Further optimization needs to be performed to improve and verify
proper adhesion prior to initiating mechanical testing.
Our results show variability in mechanical properties of MAVL
tissues derived from different mice. Balani et al. [8] report a dif-
ference in mechanical stiffness of MAVL tissues in areas of pig-
ment compared to nonpigmented areas. Mouse aortic valves have
asymmetric melanocytic pigment expression across the three leaf-
lets, which could contribute to the observed variability between
samples. We have consistently observed elevated pigmentation in
the right and noncoronary MAVLs compared to the left coronary
MAVL (Fig. 1(c)). In this study, we did not track the MAVL (i.e.,
left coronary, right coronary, or noncoronary) used from each
mouse for the tests. Future studies tracking differences between
the different MAVLs could yield insight into structural and
mechanical differences within the aortic valve.
In addition to passive responses from extracellular matrix align-
ment, the tissue response to the applied tensile load may also be
influenced by active cellular control of MAVL tonality. Dopamine
is a common vasoconstriction stimulant used to treat various car-
diovascular conditions such as low blood pressure and low heart
rate and is synthesized in the synaptic space between neurons
[17]. In this study, we coat the membrane with dopamine followed
by thorough washing prior to adding the tissue. Therefore, only
endothelial cells should contact the dopamine coating and not the
contractile cell populations that may control valve tone within the
tissue. However, future studies would need to assess the effect of
dopamine-coat on tissue biomechanics by using other adhesives
and/or blocking dopamine receptors. Decoupling these responses
in future studies will allow for a holistic understanding of the
active and passive mechanical properties of WT mouse aortic
valve tissues as well as those of genetically modified mice recapit-
ulating aspects of human valve disease progression. Tissues seem
to remain viable (Fig. 3(a)) with structurally intact extracellular
matrix components (Fig. 3(b)) after resection and mechanical test-
ing regimen. Valve interstitial cell contractility may influence the
basal tonality of AV tissue [18]. A MAVL basal tone could be
determined using this methodology in conjunction with the meth-
ods proposed by Chester et al. [19] in which inhibition and activa-
tion of smooth muscle cell contractility uncovers the role these
cell types play on AV tissue mechanics.
Acknowledgment
The authors thank Kelsy Scott and Shreyanshu Ray for techni-
cal assistance, Drs. Jorge Riera, Alexander Agoulnik, and Lidia
Kos for providing mouse tissues used in the study, and Dr. Niko-
laos Tsoukias for providing access to equipment required for
MAVL dissection.
Funding Data
Ronald E. McNair Graduate Fellowship.
A Scientist Development Grant from the American Heart
Association (17SDG633670259; Funder ID: 10.13039/
100000968).
References
[1] Sacks, M. S., and Yoganathan, A. P., 2007, “Heart Valve Function: A Biome-
chanical Perspective,” Philos. Trans. R. Soc. London, Ser. B,362(1484),
pp. 1369–1391.
[2] Tesfamariam, M. D., Mirza, A. M., Chaparro, D., Ali, A. Z., Montalvan, R.,
Saytashev, I., Gonzalez, B. A., Barreto, A., Ramella-Roman, J., Hutcheson, J.
D., and Ramaswamy, S., 2019, “Elastin-Dependent Aortic Heart Valve Leaflet
Curvature Changes During Cyclic Flexure,” Bioengineering (Basel),6(2),
p. 39.
[3] Pham, T., Sulejmani, F., Shin, E., Wang, D., and Sun, W., 2017, “Quantification
and Comparison of the Mechanical Properties of Four Human Cardiac Valves,”
Acta Biomater.,54, pp. 345–355.
[4] Stella, J. A., Liao, J., and Sacks, M. S., 2007, “Time-Dependent Biaxial
Mechanical Behavior of the Aortic Heart Valve Leaflet,” J. Biomech.,40(14),
pp. 3169–3177.
[5] Stella, J. A., and Sacks, M. S., 2007, “On the Biaxial Mechanical Properties of
the Layers of the Aortic Valve Leaflet,” ASME J. Biomech. Eng.,129(5),
pp. 757–766.
[6] Sider, K. L., Blaser, M. C., and Simmons, C. A., 2011, “Animal Models of
Calcific Aortic Valve Disease,” Int. J. Inflam.,2011, pp. 1–18.
[7] Hinton, R. B., Jr., Alfieri, C. M., Witt, S. A., Glascock, B. J., Khoury, P. R.,
Benson, D. W., and Yutzey, K. E., 2008, “Mouse Heart Valve Structure and
Function: Echocardiographic and Morphometric Analyses From the Fetus
Through the Aged Adult,” Am. J. Physiol. Heart Circ. Physiol.,294(6),
pp. H2480–H2488.
[8] Balani, K., Brito, F., Kos, L., and Agarwal, A., 2009, “Melanocyte Pigmenta-
tion Stiffens Murine Cardiac Tricuspid Valve Leaflet,” J. R. Soc. Interface,
6(40), pp. 1097–1102.
[9] Krishnamurthy, V. K., Guilak, F., Narmoneva, D. A., and Hinton, R. B., 2011,
“Regional Structure-Function Relationships in Mouse Aortic Valve Tissue,”
J. Biomech.,44(1), pp. 77–83.
[10] Sewell-Loftin, M. K., Brown, C. B., Baldwin, H. S., and Merryman, W. D.,
2012, “A Novel Technique for Quantifying Mouse Heart Valve Leaflet
Stiffness With Atomic Force Microscopy,” J. Heart Valve Dis.,21(4),
pp. 513–520.
[11] Ayoub, S., Ferrari, G., Gorman, R. C., Gorman, J. H., Schoen, F. J., and Sacks,
M. S., 2016, “Heart Valve Biomechanics and Underlying Mechanobiology,”
Compr. Physiol.,6(4), pp. 1743–1780.
[12] Yang, K., Lee, J. S., Kim, J., Lee, Y. B., Shin, H., Um, S. H., Kim, J. B., Park,
K. I., Lee, H., and Cho, S. W., 2012, “Polydopamine-Mediated Surface Modifi-
cation of Scaffold Materials for Human Neural Stem Cell Engineering,” Bioma-
terials,33(29), pp. 6952–6964.
[13] Poh, C. K., Shi, Z., Lim, T. Y., Neoh, K. G., and Wang, W., 2010, “The Effect
of VEGF Functionalization of Titanium on Endothelial Cells In Vitro,” Bioma-
terials,31(7), pp. 1578–1585.
[14] Lynge, M. E., van der Westen, R., Postma, A., and St€
adler, B., 2011,
“Polydopamine–A Nature-Inspired Polymer Coating for Biomedical Science,”
Nanoscale,3(12), pp. 4916–4928.
[15] Vande Geest, J. P., Di Martino, E. S., and Vorp, D. A., 2004, “An Analysis of
the Complete Strain Field Within Flexercell
TM
Membranes,” J. Biomech.,
37(12), pp. 1923–1928.
[16] Anssari-Benam, A., Bader, D. L., and Screen, H. R., 2011, “A
Combined Experimental and Modelling Approach to Aortic Valve
Viscoelasticity in Tensile Deformation,” J. Mater. Sci. Mater. Med.,22(2), pp.
253–262.
[17] Swann, J. W., Sinback, C. N., Pierson, M. G., and Carpe nter, D. O.,
1982, “Dopamine Produces Muscle Contractions and Modulates Motoneuron-
Induced Contractions in Aplysia Gill,” Cell. Mol. Neurobiol.,2(4), pp.
291–308.
[18] Merryman, W. D., Huang, H. Y., Schoen, F. J., and Sacks, M. S., 2006, “The
Effects of Cellular Contraction on Aortic Valve Leaflet Flexural Stiffness,”
J. Biomech.,39(1), pp. 88–96.
[19] Chester, A. H., Kershaw, J. D., Sarathchandra, P., and Yacoub, M. H., 2008,
“Localisation and Function of Nerves in the Aortic Root,” J. Mol. Cell.
Cardiol.,44(6), pp. 1045–1052.
Journal of Biomechanical Engineering OCTOBER 2020, Vol. 142 / 100801-7
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