Hyperoxia alters the mechanical properties of alveolar epithelial cells.

Abstract

Patients with severe acute lung injury are frequently administered high concentrations of oxygen (>50%) during mechanical ventilation. Long-term exposure to high levels of oxygen can cause lung injury in the absence of mechanical ventilation, but the combination of the two accelerates and increases injury. Hyperoxia causes injury to cells through the generation of excessive reactive oxygen species. However, the precise mechanisms that lead to epithelial injury and the reasons for increased injury caused by mechanical ventilation are not well understood. We hypothesized that alveolar epithelial cells (AECs) may be more susceptible to injury caused by mechanical ventilation if hyperoxia alters the mechanical properties of the cells causing them to resist deformation. To test this hypothesis, we used atomic force microscopy in the indentation mode to measure the mechanical properties of cultured AECs. Exposure of AECs to hyperoxia for 24 to 48 h caused a significant increase in the elastic modulus (a measure of resistance to deformation) of both primary rat type II AECs and a cell line of mouse AECs (MLE-12). Hyperoxia also caused remodeling of both actin and microtubules. The increase in elastic modulus was blocked by treatment with cytochalasin D. Using finite element analysis, we showed that the increase in elastic modulus can lead to increased stress near the cell perimeter in the presence of stretch. We then demonstrated that cyclic stretch of hyperoxia-treated cells caused significant cell detachment. Our results suggest that exposure to hyperoxia causes structural remodeling of AECs that leads to decreased cell deformability.

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doi: 10.1152/ajplung.00223.2011
302:L1235-L1241, 2012. First published 30 March 2012;
Am J Physiol Lung Cell Mol Physiol
Sinclair and Christopher M. Waters
Esra Roan, Kristina Wilhelm, Alex Bada, Patrudu S. Makena, Vijay K. Gorantla, Scott E.
epithelial cells
Hyperoxia alters the mechanical properties of alveolar
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Hyperoxia alters the mechanical properties of alveolar epithelial cells
Esra Roan,1Kristina Wilhelm,2Alex Bada,1Patrudu S. Makena,3Vijay K. Gorantla,3Scott E. Sinclair,2,3
and Christopher M. Waters2,3
Departments of1Biomedical Engineering and Mechanical Engineering, University of Memphis; Departments of2Physiology
and3Medicine, University of Tennessee Health Science Center, Memphis, Tennessee
Submitted 1 July 2011; accepted in final form 27 March 2012
Roan E, Wilhelm K, Bada A, Makena PS, Gorantla VK,
Sinclair SE, Waters CM. Hyperoxia alters the mechanical prop-
erties of alveolar epithelial cells. Am J Physiol Lung Cell Mol
Physiol 302: L1235–L1241, 2012. First published March 30, 2012;
doi:10.1152/ajplung.00223.2011.—Patients with severe acute lung
injury are frequently administered high concentrations of oxygen
(?50%) during mechanical ventilation. Long-term exposure to
high levels of oxygen can cause lung injury in the absence of
mechanical ventilation, but the combination of the two accelerates
and increases injury. Hyperoxia causes injury to cells through the
generation of excessive reactive oxygen species. However, the
precise mechanisms that lead to epithelial injury and the reasons
for increased injury caused by mechanical ventilation are not well
understood. We hypothesized that alveolar epithelial cells (AECs)
may be more susceptible to injury caused by mechanical ventilation
if hyperoxia alters the mechanical properties of the cells causing them to
resist deformation. To test this hypothesis, we used atomic force micros-
copy in the indentation mode to measure the mechanical properties of
cultured AECs. Exposure of AECs to hyperoxia for 24 to 48 h caused a
significant increase in the elastic modulus (a measure of resistance to
deformation) of both primary rat type II AECs and a cell line of mouse
AECs (MLE-12). Hyperoxia also caused remodeling of both actin and
microtubules. The increase in elastic modulus was blocked by treat-
ment with cytochalasin D. Using finite element analysis, we showed
that the increase in elastic modulus can lead to increased stress near
the cell perimeter in the presence of stretch. We then demonstrated
that cyclic stretch of hyperoxia-treated cells caused significant cell
detachment. Our results suggest that exposure to hyperoxia causes
structural remodeling of AECs that leads to decreased cell deform-
ability.
acute respiratory distress syndrome; atomic force microscopy; force
maps; elastic modulus
THE MORTALITY RATE OF 40–60% for patients with acute lung
injury (ALI) or acute respiratory distress syndrome (ARDS)
remains remarkably high in the United States (36). Patients
with ALI and ARDS exhibit compromised respiratory function
that is partly related to edema formation attributable to loss of
endothelial and epithelial barrier integrity (26, 40, 42). Fluid
accumulation in the alveolar space ultimately diminishes the
efficiency of gas exchange, causing hypoxemia. Although
patients are often initially treated with oxygen by mask, the
extent of injury frequently requires mechanical ventilation with
supplemental oxygen. Although mechanical ventilation is nec-
essary for the survival of patients with the most severe phys-
iological impairment, additional injury can occur, and this is
referred to as ventilator-induced lung injury (12, 23, 33). Both
mechanical ventilation and long-term exposure to hyperoxia
can independently cause lung injury that is similar histologi-
cally to that seen in ARDS (1, 10, 24). Furthermore, animal
studies have shown that the use of high levels of inspired
oxygen (FIO2?50%) can accelerate and increase lung injury
when used in conjunction with high-stretch mechanical venti-
lation (2, 10, 17, 19, 20, 31). However, the mechanisms
responsible for this enhanced injury are not well understood.
In animal studies, hyperoxia exposure for 3 days caused injury
to alveolar type I (ATI) cells, hyperproliferation of alveolar type
II (ATII) cells, diffuse hyaline membrane deposition, edema,
interstitial fibrosis, and pulmonary vascular remodeling (3, 4, 8,
15, 31). It has been proposed that hyperoxia-induced lung injury
results from increased production of reactive oxygen species,
causing either direct tissue and protein damage or activation of
signaling pathways that promote cell injury (6, 14, 16, 37). This
injury is linked to the activation of molecular pathways, leading to
both apoptosis and necrosis, but the precise pathways have not
been fully elucidated (7, 38, 44).
Previous studies have shown that oxidative stress changed
cell morphology by disrupting microfilaments and the spatial
organization of actin (9, 25, 30). Exposure of lung macro-
phages to hyperoxia caused excessive polymerization of actin,
which adversely affected the antibacterial function of the cells
(25). Endothelial cells exposed to hyperoxia exhibited an
increase in actin stress fibers (30). On the basis of these earlier
studies, we hypothesized that changes in cytoskeletal structure
caused by hyperoxia might decrease the mechanical deform-
ability of the cells and thereby enhance the development of
injury when cells are stretched during mechanical ventilation.
To test this hypothesis, we measured the elastic modulus of
cultured alveolar epithelial cells (AECs) using atomic force
microscopy (AFM) in the indentation mode. The elastic mod-
ulus, derived from direct contact of the probe with the live cell,
provides a measure of the local deformability of the cell. Cells
exposed to hyperoxia exhibited a significant increase in elastic
modulus, and this was consistent with cytoskeletal changes
observed by fluorescence microscopy. Moreover, we used a
computational approach to predict that the increase in elastic
modulus leads to increased internal cell stresses when they are
exposed to stretch, which could lead to mechanical injury of
epithelial cells. We then demonstrated that cyclic stretch of
hyperoxia-treated cells caused increased detachment of the
cells, which did not occur in control cells.
MATERIALS AND METHODS
Cell culture. Two cell types were utilized in this study: primary rat
ATII cells and a mouse alveolar epithelial cell line (MLE-12) [Amer-
ican Type Culture Collection (ATCC), Manassas, VA]. Primary rat
ATII cells were isolated according to the methods described previ-
ously (11, 43). Briefly, ATII cells were isolated from male Sprague-
Dawley rats by elastase digestion and differential adherence on
Address for reprint requests and other correspondence: C. M. Waters, Dept.
of Physiology, The Univ. of Tennessee Health Science Center, 894 Union
Ave., Rm. 426, Memphis, TN 38163-0001 (e-mail: cwaters2@uthsc.edu).
Am J Physiol Lung Cell Mol Physiol 302: L1235–L1241, 2012.
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IgG-coated dishes. ATII cells were cultured on Corning (Corning,
Corning, NY) cell culture dishes coated with a rat lung fibroblast
matrix deposited by RLF-6 cells (ATCC). Freshly isolated cells were
seeded at a density of 3.5 ? 106cells/ml in 3 ml of ATII culture
medium (DMEM; GIBCO, Carlsbad, CA) containing 10% heat-
inactivated FBS (Cell-gro, Herndon, VA, or GIBCO), 1 mM glu-
tamine, 1% penicillin/streptomycin, and 0.25 ?M amphotericin B on
60-mm cell culture dishes. Mouse alveolar epithelial MLE-12 cells
were cultured in MLE-12 culture medium (DMEM with 10% heat-
inactivated FBS, 1 mM glutamine, 1% penicillin/streptomycin). The
plating density was 1.0 ? 106cells/ml.
ATII and MLE-12 cells were divided into two groups once con-
fluent, hyperoxia and normoxia. In a 37°C incubator, confluent cells
were exposed to either room air (with 5% CO2) or to hyperoxia
(?80–90% O2, 5% CO2, balance N2) in a contained chamber for 24
h (ATII cells) or 48 h (MLE-12 cells). ATII cells were placed in the
hyperoxia chamber 3 days after cell isolation. Because ATII cells
undergo differentiation into a type I-like phenotype between 2 and 5
days in culture, and because of the potential increased sensitivity of
the primary cells, we limited the exposure to hyperoxia to 24 h. To
determine the effects of longer exposure, we examined the response of
MLE-12 cells after 48 h. To determine the effect of cytoskeletal
disruption, MLE-12 cells were treated with cytochalasin D (1 ?g/ml;
Sigma Aldrich, St. Louis, MO) for 30 min. Cells were then rinsed
three times and transferred immediately to the AFM or fixed for
microscopy.
Fluorescence staining and confocal microscopy. Following treat-
ment, monolayers were fixed in 10% formalin for 10 min and stained
simultaneously for F-actin using rhodamine-phalloidin (Cytoskeleton,
Denver, CO) and for microtubules using an antibody against ?-tubulin
conjugated with Alexa488 (Invitrogen, Carlsbad, CA) according to
the manufacturer’s protocol. For both fluorophores, a 1:250 dilution
was applied for one 1 h. Images were acquired on a Zeiss 710 (Zeiss,
Jena, Germany) laser confocal inverted microscope with six laser lines
on a ?40 objective, using the Zen 2010 software. Further image
analysis was performed with ImageJ 1.42 image processing software
(W.S. Rasband; NIH, Bethesda, MD; http://rsbweb.nih.gov/ij/).
Indentation with AFM. To measure the elastic modulus of cells, we
utilized the AFM as a nano-indenter with a flexible cantilever beam.
The AFM indentation is achieved when the cantilever is lowered
(z-displacement) enough to come in contact with the cell and cause
indentation. The force required to compress the cell structures is
related to the elastic modulus of the cell, the nominal stiffness of the
cantilever beam (its spring constant), and the geometry of the tip (see
below) (18, 32, 34). Because the cytoskeletal structure of the cell
below the indented location is heterogeneous, the resulting force-
indentation curve is an average resistance of the nearby affected
proteins, such as microtubules, actin, cortical filaments, etc.
Live cells were indented using an AFM (MFP3D; Asylum Research,
Santa Barbara, CA) with triangular silicon nitride cantilevers (SiNi;
Budget Sensors, Sofia, Bulgaria). Although the nominal stiffness of the
cantilever beams was 0.27 N/m, because of the large variation in the
actual values, the cantilever stiffness was measured at the beginning of
each experiment by generating a force-deflection (F-d) curve on a
relatively stiff substrate (the culture dish). Next, at randomly selected
spots on the confluent monolayers of cells, force-indentation curves were
recorded over a rectangle that is 3 ?m ? 50 ?m. A MATLAB (Math-
Works, Natick, MA) code that was developed in our group was used to
batch process the force-indentation curves and to determine the elastic
modulus from these curves. Using the Hertz model for a pyramidal
indenter, the elastic modulus at each location was calculated by the
following relationship E ? F [2(1 ? ?2)]/[1.4906 ?2tan(?)] (1), where E
is elastic modulus, F is force, ? is Poisson’s ratio, ? is the tip half-opening
angle, and ? is the sample indentation (18, 32, 34). In the analyses, the
Poisson’s ratio is assumed to be 0.49, which is consistent with prior
literature suggesting cells to be nearly incompressible. The indentation
depth was chosen to be 0.3 ?m to avoid the substrate stiffness concerns,
but to allow a substantial indentation to record an average local mechan-
ical response.
Initially we recorded complete maps (50 ?m by 50 ?m) with 5,184
individual data force-indentation curves. However, these required ?90
min to acquire, and we decided to record smaller maps from more
locations. To compare treatment groups, we performed measurements at
15 to 20 different locations within a culture dish with 2 ? 72 force-
indentation curves at each location. We limited the indentation depth on
the cell to 0.3 ?m corresponding to the analyzed portion of the curve. We
computed the median modulus at each location and averaged these over
the dish to obtain a measurement for that condition (over 2,000 measure-
ments for each data point).
Cell detachment. Confluent monolayers of MLE-12 cells were
exposed to either normoxia or hyperoxia for 48 h as described above
and then exposed to cyclic stretch using the Flexercell FX-4000T
tension unit (Flexcell International, Hillsborough, NC). Cells were
exposed to 20% linear strain for 60 min at a frequency of 15
cycles/min. Cells were then fixed in 10% formalin for 5 min, and
phase-contrast images were collected at ?10 magnification using an
EVOS microscope (Advanced Microscopy Group, Bothell, WA) with
three to five fields for each well. The images were then analyzed using
MATLAB software to outline areas of cell detachment and determine
the percentage of total area with denuded cells. Unstretched cells were
used as controls, and measurements of each condition were made from
three different experiments.
Statistical analysis. We utilized MATLAB and SigmaStat for the
statistical analyses. We averaged the medians of each small E-map
taken from different locations within a dish. Because of potential
variations in cells, we performed measurements of elastic modulus on
control and treated cells on the same day and performed paired
comparisons. We conducted a paired t-test by isolations or cell-
seeding events to determine whether significant differences exist
between normoxia- and hyperoxia-treated cells on a given isolation or
cell seeding events. A P value ?0.05 was considered significant.
RESULTS
Hyperoxiacausedremodelingofactinandmicrotubulestructures.
To examine whether hyperoxia caused remodeling of the
cytoskeleton through reorganization of actin and microtubules,
we exposed ATII and MLE-12 cells to either normoxia or
80–90% O2and then stained for F-actin and ?-tubulin. Rep-
resentative images of ATII and MLE-12 cells treated with
normoxia or hyperoxia are shown in Fig. 1. In control ATII
cells, we observed typical F-actin staining with thin filaments
both in the cortical regions and in the central area of the cell
(Fig. 1A). Following exposure to hyperoxia, larger and thicker
F-actin bands were observed both in the cell interior and in
cortical regions (Fig. 1C). Microtubules appeared to be more
densely distributed in hyperoxia-treated ATII cells and some-
what restricted from the cortical regions (Fig. 1, B and D). In
control MLE-12 cells, there were fewer F-actin filaments
spanning the cell compared with ATII cells, in part due to the
prominent nuclei (not shown in these images), but there was
strong cortical staining (Fig. 1E). Hyperoxia caused an increase
in both cortical actin and F-actin filaments in MLE-12 cells
(Fig. 1G). There was also stronger cortical staining of micro-
tubules in hyperoxia-treated MLE-12 cells (Fig. 1, F and H). In
both cell types, there was some nuclear damage following
hyperoxia treatment.
Hyperoxia increased the elastic modulus of alveolar epithe-
lial cells. Because hyperoxia caused significant changes in
actin and microtubule distribution, we hypothesized that this
would also lead to changes in the elastic moduli. To test this,
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we exposed ATII and MLE-12 cells to either normoxia or
80–90% O2and then measured the elastic modulus. Examples
of detailed E-maps for normoxia- and hyperoxia-treated
MLE-12 cells are shown in Fig. 2. These maps illustrate the
variability of elastic modulus within a given field of cells and
show that hyperoxia treatment caused more sights of increased
stiffness. As shown in Fig. 2C, hyperoxia caused a significant
increase in modulus for both ATII and MLE-12 cells, indicat-
ing a decrease in the deformability of the cells.
Cytochalasin D reduced the elastic modulus. To determine
the extent to which the mechanical response that we measured
(elastic modulus) was dependent on the actin cytoskeleton, we
measured the elastic modulus of MLE-12 cells following
treatment with either normoxia or hyperoxia followed by
treatment with cytochalasin D (cytoD) to disrupt F-actin. As
shown in Fig. 1, I–J, cytoD treatment caused disruption of
F-actin as well as microtubules in normoxia-treated cells.
CytoD treatment caused similar disruption in hyperoxia-treated
cells (not shown). Importantly, cytoD caused a significant
reduction in elastic modulus in both normoxia- and hyperoxia-
treated MLE-12 cells (Fig. 3).
Finite element analysis predicts higher internal stress near
the cell edge in hyperoxia-treated cells. The increase in elastic
modulus of the cells could lead to a modification of the
stress-strain profiles experienced by cells exposed to injurious
levels of distention. To investigate whether the decrease in cell
deformability caused by hyperoxia would alter the response of
the cell to a large deforming stress, we developed a finite
element model of a cell residing on a flexible substrate. We
utilized finite element analysis (ABAQUS; Simulia, Provi-
dence, RI) to show how internal stresses would be altered when
a cell was exposed to quasistatic injurious stretch applied in the
plane of a flexible substrate. As shown in Fig. 4A, we generated
a two-dimensional (2D) axis-symmetric model (right) of a cell
(gray) residing on a flexible membrane (blue), which is me-
chanically equivalent to the three-dimensional system shown
on the left. We conducted two different analyses in which the
membrane had an elastic modulus of E ? 5.0 kPa (5) in both
cases, and the modulus of the cell, Ecell, was modified accord-
ing to our experimental findings of normoxia- and hyperoxia-
treated primary ATII cells. A significant increase in the cell
elastic modulus could lead to elevated stress levels because of
increased dissimilarity between the cell elastic modulus and
that of the alveolar wall on which the cells reside.
The results of this analysis (Fig. 4C) show that a 20%
deforming stretch of the membrane would cause higher internal
stresses in the hyperoxic cells compared with the control cells
(indicated by the warmer colors in the map). For example, the
stress in the thinner regions of the cell near the perimeter is
higher in the hyperoxia cell compared with the normoxia cell.
The cell is likely experiencing a more complex stress field near
the perimeter, which is not shown for simplicity.
Cyclic stretch caused detachment of hyperoxia-treated cells.
To test the hypothesis that hyperoxia-treated cells are more
Fig. 1. Hyperoxia treatment caused changes in the distribution of F-actin (red,
A, C, E, G) and microtubules (green, B, D, F, H) in alveolar type II (ATII) cells
(A–D) and MLE-12 cells (E–H). Treatment with cytochalasin D (cytoD)
caused disruption of actin and microtubules in MLE-12 cells (I–J). Each pair
of images was from the same field. Scale bars are 50 ?m (n ? 3).
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susceptible to stretch-induced injury, we exposed MLE-12
cells to either normoxia or hyperoxia for 48 h, followed by
cyclic stretch (20%, 15 cycles/min) for 60 min and then
measured cell detachment. As shown in Fig. 5, there was little
or no cell detachment when normoxia-treated cells were ex-
posed to stretch, but there was significant cell detachment
following stretch of hyperoxia-treated cells (significantly dif-
ferent from control, P ? 0.05; n ? 3).
DISCUSSION
In this study, we provide novel data demonstrating a signif-
icant increase in the elastic modulus of cells following expo-
sure to hyperoxia that correlated with cytoskeletal remodeling.
Qualitative analyses of fluorescence images suggest that, al-
though ATII and MLE-12 cells have different structural ap-
pearance under control conditions, hyperoxia caused cytoskel-
etal remodeling in both. ATII cells exposed to hyperoxia
contain F-actin that is in short but thick bundles, indicating a
stressed condition of the whole cell. This finding is consistent
with other studies that showed stress fiber formation or exces-
sive polymerization of actin attributable to extended treatment
with high levels of oxygen (9, 25, 30). In MLE-12 cells the
changes in F-actin and microtubules were more apparent in the
cortical regions, perhaps attributable to the presence of a
relatively large nucleus in these cells (not shown). We are not
aware of previous studies demonstrating changes in microtu-
bules in response to hyperoxia. Whereas previous studies have
indicated the role of F-actin and microtubules in contributing to
the stiffness of cells by AFM (28, 35), we have shown for the
first time that hyperoxia-induced changes in actin and micro-
tubules are consistent with increased cell stiffness in two types
of cells.
As the elastic modulus of a cell increases, it becomes more
resistant to deformation. The consequences of such an increase in
modulus in lung cells may be significant given that these cells
undergo cyclic deformation during the breathing cycle (27, 29,
39). It has been estimated that the epithelial cells that reside on
the lung basement membrane undergo a tensile strain that is
?4% during normal tidal breathing (13, 27). During mechan-
ical ventilation of patients with ARDS, there may be substan-
tial heterogeneity of strain levels in the alveoli attributable to
the presence of edema fluid, overinflation of air-filled regions,
and collapse of airspaces (22, 26, 41). If mechanical ventilation
with high levels of inspired oxygen causes increased resistance
Fig. 2. Detailed E-maps of a confluent mono-
layer of MLE-12 cells treated with normoxia
(A) and hyperoxia (B). In these detailed
maps, each pixel corresponds to a single
force-indentation curve. C: elastic moduli
(E) of ATII and MLE-12 cells incubated
with normoxia or hyperoxia. ATII cells were
exposed to either normoxia or hyperoxia for
24 h on the third day after isolation. MLE-12
cells were exposed for 48 h. Hyperoxia
caused a significant increase in the elastic
modulus in each type of cell (*significantly
different from normoxia, P ? 0.05; paired
t-test; error bars indicate SE). Each bar cor-
responds to 4 separate isolations in the case
of ATII cells (n ? 4) and 5 different cell-
seeding events in the case of MLE-12 cells
(n ? 5). Each data point represents the mean
value from 15 to 20 locations in a given dish
in which the median value was determined
from 144 measurements near that location.
Fig. 3. Elastic moduli of MLE-12 cells incubated with normoxia or hyperoxia
(as described in Fig. 2) followed by treatment with cytochalasin D (cytD) for
30 min. Each bar corresponds to 3 different cell-seeding events (n ? 3) with
each data point corresponding to the mean value of the median from ?144
measurements from 15–20 different locations (*significantly different from
normoxia, P ? 0.05; error bars represent SE).
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to deformation of AECs in overdistended alveoli, this could
result in increased injury.
To reduce cell injury, the basement membrane and cells
should deform, with nearly identical intrinsic mechanical re-
sponses. This similarity of mechanical responses limits the
magnitude of cytoskeletal stresses from reaching excessive
levels and causing adverse effects such as damage to focal
adhesions or cell-to-cell junctions. Our measurements in iso-
lated cells predict that hyperoxia would cause an imbalance
between the elastic modulus of the cell and the alveolar walls.
In such a case the cellular stresses resulting from an expansion
of the basement membrane should increase as a function of the
extent of the dissimilarity of the materials. Our finite element
analysis supports this and shows that the dissimilarity induces
higher levels of stress near the perimeter of cells. Under such
conditions, the cell cytoskeleton may remodel to adjust to the
deforming stress, but we have not yet investigated this possi-
bility.
To test whether the increase in interfacial stress between the
cell and the substrate could lead to injury, we mechanically
stretched cells that were exposed to hyperoxia. Although we
found little detachment of control, normoxia-treated cells,
stretch of hyperoxia-treated cells caused significant cell detach-
ment (Fig. 5). According to the finite element analysis, the
most likely explanation for the formation of gaps in the
monolayer would be the disruption of cell-substrate connec-
Fig. 4. A: 3D depiction of the cell on the
flexible membrane. B: 2D finite element model
of the cell-membrane system where a lateral
stretch is applied on the membrane. C: contour
plots of the stress in the direction parallel to the
20% stretch load showing an increased tensile
stress (?x) in the hyperoxia-treated cells near
the cell boundary.
Fig. 5. Hyperoxia treatment followed by cy-
clic stretch caused detachment of MLE-12
cells. Cells were treated for 48 h with nor-
moxia (A and C) or hyperoxia (B and D) and
then kept static (A and B) or exposed to 20%
cyclic stretch, 15 cycles/min (C and D) for
60 min. E: percentage of area of detached
cells was determined for at least 5 fields
from at least 3 different wells (n ? 3; *sig-
nificantly different from unstretched nor-
moxia, P ? 0.05).
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tions. This is due to the high in-plane stress at the cell
periphery, which would apply a combined shear and tensile
stress on the cell-substrate junctions. However, other potential
forms of injury such as plasma membrane wounding are also
possible.
There are other limitations to our study. The amount of time
required to perform AFM indentation measurements limits our
ability to measure maps with high spatial resolution while
sampling a sufficient number of cells. Because of this limita-
tion, we chose to conduct more measurements of smaller fields
(15–20 locations) rather than obtain high spatial resolution
maps (as shown in Fig. 2). This allowed us to contact a greater
number of cells with a given monolayer. Considering that we
recorded a minimum of 15 E-maps that spanned 50 ?m, it is
likely that we contacted 50 or more nonadjacent cells.
Another potential limitation is the possibility that measure-
ments may be affected by the underlying substrate. Because
AFM indentation causes compression of the cells that are on a
relatively hard substrate, there is a concern that, if the cells are
too thin, then the measurements may be affected by the
presence of the substrate. To address this concern, we con-
firmed that the cells were tall enough to be indented an optimal
0.3 ?m. Because of the chosen indentation depth, our mea-
surements are less likely to be affected by the nucleus. In our
E-maps we did not observe a high elastic modulus because of
the presence of the nucleus, which is known to be higher than
the cell cytoplasm (21).
In this study, we assumed that the cell was nearly incom-
pressible, i.e., a Poisson’s ratio of 0.49. Although cells are
mostly composed of incompressible fluid, there may exist local
fluctuations in Poisson’s ratio. However, it is unlikely that such
a spatial variation in Poisson’s ratio would account for the
significant changes in elastic modulus that we observed in
hyperoxia-treated cells. The Poisson’s ratio for an object com-
posed of mostly fluid would be unlikely to fall below 0.45, and
such a change would not account for the differences that we
observed.
The elastic modulus of control ATII cells was significantly
higher than that of control MLE-12 cells. MLE-12 cells were
relatively taller than the ATII cells (data not shown), and
F-actin fibers located centrally in the primary cells were not as
evident in the MLE-12 cells. As indicated by comparing Figs.
1 and 2, there appear to be more prominent increases in elastic
modulus in MLE-12 cells in what appears to be the cell-cell
junctions, but we cannot confirm this without simultaneous
imaging of the cells with force map measurements. In ATII
cells, the changes appear to occur throughout the cells. Nev-
ertheless, the fold change in the elastic modulus of hyperoxic
over the normoxic cells was nearly the same for primary and
MLE-12 cells, 1.42 and 1.63, respectively. Treatment of MLE-12
cells with cytoD caused a significant decrease in the elastic
modulus, suggesting that F-actin contributes to the mechanical
response of these cells to indentation. In addition, cytoD caused a
reduction in elastic modulus in hyperoxia-treated cells to the level
of normoxia-treated cells, suggesting that the changes in elastic
modulus with hyperoxia were attributable to changes in actin.
We should note here that we exposed the MLE-12 cells to
hyperoxia for 48 h, whereas ATII cells were exposed for 24 h.
As mentioned in MATERIALS AND METHODS, we wanted to limit
the time of exposure of ATII cells to minimize the effects of
differentiation of ATII cells into ATI cells during the experi-
ment. In future studies, we plan to investigate the development
of the changes in mechanical properties with time of exposure
to hyperoxia.
Our finite element analysis is limited as a result of simpli-
fications, but it is useful in illustrating the principles of how
stresses may be altered in cells when there are differences in
material properties between the cells and the substrate. Al-
though this model oversimplifies the in vivo conditions, which
are likely to include stresses in multiple directions and trans-
mission of forces attributable to cell-cell contacts, our analysis
suggests that stretch induces stresses near the cell borders. Our
analysis also ignores the potential for changes in the mechan-
ical properties of the extracellular matrix on which the cells are
attached. It is possible that such changes also occur with
hyperoxia, but there are likely differences in the time course of
such events with cytoskeletal changes likely to occur sooner.
Our simplistic analysis indicating increased stress within cells
during stretch is supported by our experimental results dem-
onstrating detachment of hyperoxia-treated cells following
stretch (Fig. 5).
In summary, we utilized AFM indentation to show that the
elastic modulus of live alveolar epithelial cells was increased
following exposure to hyperoxia. We provided additional qual-
itative evidence showing the remodeling of ATII and MLE-12
cells showing significant alterations in F-actin and microtu-
bules. Hyperoxia-treated cells showed increased detachment
when subsequently exposed to mechanical stretch. Our find-
ings regarding the changes in elastic modulus in conjunction
with finite element modeling suggest a potential mechanism for
enhanced epithelial injury during mechanical ventilation with
hyperoxia.
ACKNOWLEDGMENTS
We thank Kathy Troughten at the University of Tennessee Health Science
Center for help in confocal microscopy. We are grateful to Dr. Manik Ghosh,
Charlean L. Luellen, and Dr. Lynn M. Crosby at the University of Tennessee
Health Science Center for help, advice, and technical assistance. We thank Dr.
Lindner and Dr. Pinkhasshik at the University of Memphis for the unlimited
access to the AFM.
GRANTS
This work was supported by NIH grant HL094366.
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the authors.
AUTHOR CONTRIBUTIONS
Author contributions: E.R., K.W., A.B., P.S.M., V.K.G., S.E.S., and
C.M.W. conception and design of research; E.R., K.W., A.B., P.S.M., V.K.G.,
and C.M.W. performed experiments; E.R., K.W., A.B., P.S.M., and C.M.W.
analyzed data; E.R., K.W., P.S.M., V.K.G., S.E.S., and C.M.W. interpreted
results of experiments; E.R., K.W., and C.M.W. prepared Figs.; E.R., K.W.,
and C.M.W. drafted manuscript; E.R., K.W., P.S.M., S.E.S., and C.M.W.
edited and revised manuscript; E.R., K.W., and C.M.W. approved final version
of manuscript.
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