Am J Physiol Lung Cell Mol Physiol 306: L672–L683, 2014. First published January 31, 2014; doi:10.1152/ajplung.00106.2013.

Microfluidic shear stress-regulated surfactant secretion in alveolar epithelial type II cells in vitro Sanjeev Kumar Mahto,1 Janna Tenenbaum-Katan,1 Ayala Greenblum,1 Barbara Rothen-Rutishauser,2 and Josué Sznitman1 1

Department of Biomedical Engineering, Technion-Israel Institute of Technology, Haifa, Israel; and 2Adolphe Merkle Institute, University of Fribourg, Marly, Switzerland Submitted 24 April 2013; accepted in final form 23 January 2014

Mahto SK, Tenenbaum-Katan J, Greenblum A, Rothen-Rutishauser B, Sznitman J. Microfluidic shear stress-regulated surfactant secretion in alveolar epithelial type II cells in vitro. Am J Physiol Lung Cell Mol Physiol 306: L672–L683, 2014. First published January 31, 2014; doi:10.1152/ajplung.00106.2013.—We investigated the role of flow-induced shear stress on the mechanisms regulating surfactant secretion in type II alveolar epithelial cells (ATII) using microfluidic models. Following flow stimulation spanning a range of wall shear stress (WSS) magnitudes, monolayers of ATII (MLE-12 and A549) cells were examined for surfactant secretion by evaluating essential steps of the process, including relative changes in the number of fusion events of lamellar bodies (LBs) with the plasma membrane (PM) and intracellular redistribution of LBs. F-actin cytoskeleton and calcium levels were analyzed in A549 cells subjected to WSS spanning 4 –20 dyn/cm2. Results reveal an enhancement in LB fusion events with the PM in MLE-12 cells upon flow stimulation, whereas A549 cells exhibit no foreseeable changes in the monitored number of fusion events for WSS levels ranging up to a threshold of ⬃8 dyn/cm2; above this threshold, we witness instead a decrease in LB fusion events in A549 cells. However, patterns of LB redistribution suggest that WSS can potentially serve as a stimulus for A549 cells to trigger the intracellular transport of LBs toward the cell periphery. This observation is accompanied by a fragmentation of F-actin, indicating that disorganization of the F-actin cytoskeleton might act as a limiting factor for LB fusion events. Moreover, we note a rise in cytosolic calcium ([Ca2⫹]c) levels following stimulation of A549 cells with WSS magnitudes ranging near or above the experimental threshold. Overall, WSS stimulation can influence key components of molecular machinery for regulated surfactant secretion in ATII cells in vitro. alveolar epithelial cells; pulmonary surfactant; lamellar bodies; Factin cytoskeleton; shear stress; microfluidics

(ATII) are mainly involved in the synthesis and secretion of pulmonary surfactant in the human lung. Under physiological conditions, mechanical stretching resulting from parenchymal wall distensions during breathing is considered a primary stimulus for surfactant secretion in ATII cells (9, 14, 15, 30). Pulmonary surfactant is composed of ⬃90% lipids, mainly saturated phosphatidylcholine, and ⬃10% proteins, including the surfactant apoproteins SP-A, SP-B, SP-C, and SP-D (31, 38). These cells contain characteristic organelles, often referred to as “lamellar bodies” (LBs) that store surfactant phospholipids and a variety of proteins (2). LBs secrete their contents into the alveolar space via exocytosis that is considered a slow, complex, and multi-

TYPE II ALVEOLAR EPITHELIAL CELLS

Address for reprint requests and other correspondence: J. Sznitman, Technion - Israel Institute of Technology, Haifa, Israel (e-mail: sznitman@bm. technion.ac.il). L672

phasic process and is thought to be initiated by a number of chemical and physical stimuli (2, 22, 52). Two noteworthy steps are involved in the final stage of the secretory process and include 1) the translocation of LBs to the apical plasma membrane (PM), followed by 2) the fusion of LBs with the PM (33, 43). Previous studies have also confirmed that cortical cytoskeleton (actin) plays an important role in regulating exocytosis of LBs (33, 41). Similar to the general mechanism of exocytosis, substantial direct and indirect evidence suggests that cytosolic calcium ([Ca2⫹]c) acts as a second messenger for LB exocytosis (5, 6). Namely, a growing body of work has suggested that both chemical and physical stimuli-induced LB exocytosis is regulated by calcium (18, 23). Although the role of mechanical stretching stimuli on regulated surfactant secretion in ATII cells has been increasingly characterized (2, 3, 12, 44), the influence of fluid flow-induced shear stresses remains still largely unknown. This latter effect is potentially relevant under various pulmonary diseases and clinical therapies, including mechanical ventilation and surfactant replacement therapy. For such physiological scenarios, liquid plugs that form may potentially occlude the air-filled lung, propagate through airways upon lung inflation during breathing, and produce localized, yet severe, fluid wall shear stresses (WSS) on the underlying epithelial monolayer, resulting in, among other things, cell damage and injury (7, 16, 25, 57). Recent simulations have estimated WSS magnitudes of several tens of dynes per square centimeter, with local maxima reaching well above 40 dyn/cm2, for the advancing and receding motion of propagating liquid plugs (16, 25). In recent years, microfluidic devices have been increasingly utilized to recreate in vitro models of cellular lung airway and alveolar environments and investigate among other cell injury under mechanical stretching conditions (16, 25, 26, 31). Here, we have tested the hypothesis of whether flow-induced shear stresses may influence the LB-dependent pathways regulating surfactant secretion in ATII cells. Specifically, MLE-12 and A549 cells, mouse respiratory and human alveolar type II like cell lines (29, 55) that are well characterized and widely used as in vitro lung epithelial cell models (42), were cultured to subconfluence inside microfluidic channels and examined for surfactant secretion under fluid shear stress stimulation spanning a range of low to high WSS magnitudes. Our experimental in vitro findings indicate that flow-induced shear stress can significantly enhance the number of fusion events of LBs with the PM for MLE-12 cells. Concurrently, fusion events of LBs in A549 cells are significantly hindered for sufficiently high WSS magnitudes, where shear stress appears to act as a stimulus for the induction of intracellular LB transport in such cells. These latter observations are found to be coupled to the

1040-0605/14 Copyright © 2014 the American Physiological Society

http://www.ajplung.org

L673

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

disorganization of F-actin cytoskeleton and calcium levels in A549 cells, where fragmentation of actin filaments might be acting as a limiting factor for LB fusion events. Altogether, our study provides important insights on the role of fluid flowinduced shear stress in regulating the process of surfactant secretion. MATERIALS AND METHODS

Fabrication of microfluidic channels. Microfluidic devices (Fig. 1A) consisting of straight channels (e.g., 0.4 mm wide, 4.5 mm long, 0.08 mm high) were fabricated in poly(dimethylsiloxane), PDMS (Sylgard 184; Dow Corning), using rapid prototyping technique and soft-lithography microfabrication, according to established procedures (32, 35). Briefly, the master for the cell culture device was fabricated by patterning negative photoresist (SU-8 2150; Microchem) on a 2⬙ silicon wafer using a mask aligner. After development, the wafer was placed in a clean Petri dish and was treated with (tridecafluoro-1,1,2,2-tetrahydrooctyl)trichlorosilane (Aldrich) to allow easy removal of the PDMS from the master mold. The mixture of PDMS, prepolymer and catalyst (10:1 ratio), was poured over the master and the Petri dish containing the master and PDMS was then placed in a dry oven for 4 h at 70°C. The inlet and outlet for the fluids were punched out using biopsy punches (Kai Medical). The surfaces of the PDMS replica and a glass substrate (i.e., glass coverslips) were activated with oxygen plasma, using a hand-held laboratory corona treater (Electro-Technic Products) and brought together by visual alignment immediately after activation to form an irreversible bond. Glass coverslips (24 ⫻ 60 mm2, no. 1, De-Groot Laboratory Equipment) were first cleaned by immersion in 2% of aqueous Micro-90 cleaning solution (Cole Parmer) followed by sonication for 2 h at room temperature. The cleaned coverslips were repeatedly rinsed in deionized water (3 times) and dried before irreversible bonding with PDMS channels (see Fig. 1A). Culture and microfluidic fluid flow stimulation of ATII cells. Two distinct ATII cell lines were investigated. Mouse lung epithelial cells (MLE-12) (American Type Culture Collection, ATCC, Biological Industries) were cultured in Dulbecco’s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F-12) (ATCC) supplemented with 10% fetal bovine serum (FBS, Biological Industries) in a humidified incubator (Thermo Scientific) at 37°C with 5% CO2 atmosphere. Human type II alveolar epithelial cells (A549) (American Type Culture Collection, ATCC, Biological Industries) were cultured in RPMI-1640 medium with L-glutamine (Sigma) supplemented with 10% FBS. Cells were routinely cultured in 100-mm Petri dishes. For microfluidic cultures, devices were first treated with 0.01% collagen (from rat tail, type 1, Sigma) for 2 h at 37°C followed by washing with PBS (Sigma) (3 times). The device was then dried and kept under ultraviolet exposure for 1 h before use. Cells were cultured within the microfluidic channels at a seeding density of 1 ⫻ 106 cells/ml (20 ␮l) suspension. For the analysis of intracellular LB distribution, fusion

A

inlet

PDMS channel

coverslip

events, and F-actin cytoskeleton, cells were permitted to grow in the devices under static conditions until they reached ⬃70 –90% confluency (Fig. 1B). For the staining of intracellular calcium, cells were cultured to slightly lower levels of confluency to precisely analyze the intensity profile of single cells. To create fluid flow-induced shear stresses, stimulations were performed using a Hanks’ Balanced Salt Solution (HBSS) medium, with calcium (Ca2⫹) and magnesium (Mg2⫹) (Biological Industries). Fluidic connections were established using polyethylene tubing (New Biotechnology) and a syringe pump (Harvard Apparatus), where we limit our experimental flow conditions to steady unidirectional flows within the devices. Here, we have investigated variable perfusion rates (i.e., 20, 40, 60, and 100 ␮l/min) compared with static conditions. These selected flow rates correspond to estimated WSS values of ⬃4, 8, 12, and 20 dyn/cm2, respectively, following the widely used expression for laminar three-dimensional flow in a rectangular channel (11, 39), WSS ⫽ 6 ␮Q/wh2, where ␮ is the dynamic viscosity of fluid, Q is the volumetric flow rate, and w and h are the flow channel width and height, respectively. Viability assay. The viability of cells under fluid shear stress conditions was determined by a combination of two fluorescent dyes, following established protocols (46): calcein acetoxymethyl (AM) (5 ␮M) and propidium iodide (PI) (5 ␮M) (AAT Bioquest). Calcein AM (green in color) and PI (red in color) were used to detect live and dead cells, respectively. After flow stimulation for 20 min at maximal perfusion rate (100 ␮l/min), cells were stained with the fluorescent dyes dissolved in HBSS medium containing probenecid (2.5 mM; Sigma) and pluronic F-127 (0.02%; Sigma) for 40 min at 37°C. Probenecid and pluronic F-127 were used to reduce leakage of the deesterified indicators and to increase the aqueous solubility of AM esters, respectively. For the negative control, 70% ethanol was used to treat the cells. At least three independent experiments were performed in each case. Fluorescence staining of phospholipids and LBs. To monitor LB fusion events with the PM, cells subjected to flow stimulation for 20 min were incubated with a cell-impermeant lipophilic fluorescent dye, MM 1-43 (4 ␮M) (AAT Bioquest) (an analog of FM 1– 43; FM is the trademark of Molecular Probes) for 3 min at 37°C temperature (Fig. 1C). To observe changes in the intracellular spatial redistribution of LBs upon flow stimulation, cells cultured in microfluidic channels were initially loaded with 1 ␮M quinacrine (Sigma), a fluorescent marker of LBs for 40 min at 37°C temperature. Fluorescent images were acquired across the microfluidic channel length to observe the distribution of LBs within the cells at t ⫽ 0. Thereafter, cells were subjected to flow stimulation for 20 min followed by the acquisition of an additional set of fluorescent images at t ⫽ 20 min (Fig. 1C). At least five independent experiments were performed in each case. Fluorescent staining of F-actin cytoskeleton. After 20 min of flow stimulation, cells inside microfluidic devices were fixed in 4% paraformaldehyde (Aldrich) for 15 min at 37°C followed by washing with PBS. Subsequently, cells were permeabilized with 0.5% Triton X-100

B

C

outlet 100 m

10 m

Fig. 1. A: schematic of a (PDMS)-based microfluidic device consisting of a straight channel (0.4 mm wide, 4.5 mm long, 0.08 mm high). B: phase-contrast image shows a monolayer of type II alveolar epithelial (A549) cells cultured inside the microfluidic channel (⬎80% confluency). C: A549 cells were subjected to a flow rate of 20 ␮l/min (corresponding to ⬃4 dyn/cm2) for 20 min and thereafter stained with fluorescent dyes, quinacrine (green) and MM 1– 43 (red), for the detection of lamellar bodies (LBs) and fusion of LBs with the plasma membrane, respectively. AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L674

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

(Aldrich) in PBS for 5 min at 4°C. Cells were then exposed to rhodamine-conjugated phalloidin (50 ␮g/ml, Sigma) for 40 min at room temperature. Samples were mounted in a FluoromountG, slide mountant (EMS) and then viewed under an inverted epifluorescence microscopy (Nikon Ti-E). Calcium staining. To monitor changes in the amount of [Ca2⫹]c upon flow stimulation, cells were first loaded with 4 ␮M Quest Fluo-8 (AAT Bioquest) for 45 min at 37°C temperature. The fluorescent dye was dissolved in HBSS medium (without Ca2⫹ and Mg2⫹) containing probenecid (2.5 mM) and pluronic F-127 (0.02%). After the loading, microfluidic channels were washed with HBSS (without Ca2⫹ and Mg2⫹) containing 2.5 mM probenecid (2 times) followed by the establishment of the fluidic connections. Under microscopic observation, fluorescence intensity was initially analyzed as a baseline for 1 min, and thereafter flow stimulation was performed for 4 min to stimulate the cells using HBSS (with Ca2⫹ and Mg2⫹). Fluorescent images were recorded using a Neo sCMOS camera (Andor Technology) with an acquisition rate of five frames per second. Note that, for such calcium recordings, we identified the best appropriate focal plane during microscopy imaging and maintained it fixed across the entire sample. Microscopy and live cell imaging system. The microfluidic device containing lung epithelial cells was placed inside an environmental device (37°C and 5% CO2) (Okolab Incubator, Agentek) on the inverted epifluorescence microscope. The microfluidic device was maintained under a constant perfusion rate of medium (see values described earlier), at a physiological pH and temperature. Representative fluorescent images and time-lapse phase-contrast images were acquired every 5 min for an hour using a ⫻20 (1.5 ⫻ zoom) objective and a mercury lamp (Nikon Intensilight C-HGFIE) illuminator. Image analysis. To estimate the number of fusion events coinciding with phospholipid release in a region of interest (ROI) containing individual cells, fluorescent images were analyzed via ImageJ software to count the average number (per cell) of MM 1– 43-stained spots at t ⫽ 20 min (Fig. 2A). Briefly, the image-processing method extracts the local maxima of luminance (i.e., grayscale intensity) in an ROI, thus providing a count of the number of maxima present. Results are presented as an average (mean) number of fusion events per cell (and corresponding standard deviation), similar to previous approaches (17, 23). The main advantage of this method is that raw images can be directly used, where an intensity threshold can be set to estimate the number of stained spots above the noise tolerance level, significantly avoiding the interference from background fluorescence (28). An additional advantage lies in that this method counts lighter as well as brighter spots as a single fusion event and eliminates potential bias resulting from the close proximity of fusion events and differences in intensity (Fig. 2A). Alternatively, the major limitation of the implemented method lies in that the number of fusion events may not be accurate because fusion events lying very close to each other cannot be differentiated easily (28). Relative changes in the spatial redistribution of intracellular LBs as a result of flow stimulation were estimated using digital imageprocessing techniques. First, a specific ROI comprising a variable number of cells (⬃1 to 5) was selected from the original backgroundcorrected grayscale images captured at two different time intervals, namely at time t ⫽ 0 (ROIt⫽0) and t ⫽ 20 min (ROIt⫽20), as depicted in Fig. 2B (top and middle). These ROIs were then binarized following selection of an appropriate threshold value, given the grayscale intensity distribution in the original ROIs (Fig. 2B, bottom, left, and middle). By subtracting the resulting images, a subtracted binary image (ROIt⫽0-ROIt⫽20) was obtained (Fig. 2B; bottom, right), capturing relative changes in the intracellular redistribution patterns associated with the translocation of LBs over a time interval. Namely, the number of binary pixels corresponding to these LB redistributions (i.e., I0-I20, where I is the number of binary pixels at time t) was counted such that the relative number (in percentage) of incidences representing intracellular shifts in LB positions over the measured

time interval is given as (I0-I20)/I0 ⫻ 100. It is important to note that the implemented method estimates neither the distance travelled by LBs nor the specific direction of movement; rather it solely quantifies the relative incidence of intracellular LB redistribution following flow stimulation. Considering the existing knowledge on the process of LB-dependent surfactant secretion (2, 12, 27, 52) and our experimental observations (Fig. 2B, middle), it may be inferred that such displacements, or drifts, in LBs as a result of flow stimulation might be interpreted as belonging to the noteworthy, final stages of the secretory process. Statistical analysis. Results are presented as means ⫾ SD. The statistical significance was determined by one-way ANOVA followed by a Tukey’s HSD post hoc test. The differences were considered statistically significant when P ⬍ 0.01. RESULTS

Characterization of microfluidic cell culture and viability. Following established morphometric measurements of alveolated airways (21, 53, 54), we have developed a simple microfluidic device consisting of straight channels of 400 ␮m in width (Fig. 1A). This specific width was chosen to avoid the limitations of margin values estimated by Weibel and Gomez (53) and to recreate in vitro the physiological scale of ATII cell environments. Microfluidic devices were successfully used for culturing MLE-12 and A549 cells and developing a monolayer (⬎80% confluency) of epithelial cells (Fig. 1B), under static conditions. A final time interval of 20 min was chosen on the basis that the morphology of cells after exposure to fluid WSS remained nearly unchanged for each experimental flow stimulation condition (i.e., 20, 40, 60, and 100 ␮l/min). A549 cells subjected to a flow rate of 40 ␮l/min (or more) for over 30 min resulted in significant alterations in morphologies, including shrinkage and round- and oval-shaped appearances (Fig. 3A). Using an image cross-correlation algorithm (Matlab), we quantitatively assessed a marked decline in the normalized correlation index (0.802 ⫾ 0.050; n ⫽ 10 pairs) for the flow condition 40 ␮l/min after 40 min, compared with those of static (0.920 ⫾ 0.034; n ⫽ 10 pairs) and 20 ␮l/min (0.904 ⫾ 0.033; n ⫽ 10 pairs) flow conditions; here, an ideal normalized correlation index of 1 would indicate that an image at time 0 and subsequently at 40 min is perfectly identical (see Appendix A). In contrast, we obtained high normalized correlation indices for measurements after 20 min across all flow conditions, i.e., 0.925 ⫾ 0.032 (n ⫽ 73 cells), 0.905 ⫾ 0.040 (n ⫽ 83 cells), 0.903 ⫾ 0.049 (n ⫽ 81 cells), 0.918 ⫾ 0.032 (n ⫽ 79 cells), and 0.916 ⫾ 0.033 (n ⫽ 70 cells) for 0, 20, 40, 60, and 100 ␮l/min, respectively. These latter results suggest that changes in cell morphology over the selected time interval (i.e., 20 min) were indeed negligible for all perfusion rates investigated. In addition, both cell types were tested for viability under the applied flow conditions, with no significant loss in viability observed when cells were subjected to a maximal flow rate of 100 ␮l/min for 20 min (Fig. 3B). In contrast, cells treated with 70% ethanol for 5 min showed a severe loss of viability with evidence of compromised PM (data not shown). Overall, our findings indicate that ATII cells were viable under the range of flow conditions tested. Effects of shear stress on LB exocytosis and distribution. Previous studies have suggested that surfactant secretion is a slow process, where the release of phospholipids is significantly delayed compared with LB fusion with the PM (14, 23, 52). Therefore, we aimed to visualize the LB contents imme-

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L675

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

A

phase-contrast

MM 1-43 staining

number of fusion events

10 m

B

t=0

t=20 min

10 m 60 L/min flow condition (12 dyne/cm2)

10 m

I0

I20

I0-I20

10 m

Fig. 2. A: method for estimating the number of fusion events per cell. Left: phase contrast image. Middle: average number of MM 1– 43-stained spots per cell was quantified via measuring the local maxima of luminance (i.e., intensity) in a region of interest (ROI). Right: extracted points yield an estimate of the number of fusion events in the ROI (see Image analysis for further details). B: image processing method to quantify the relative occurrence of LBs translocation over time. An example is shown for phase-contrast images (top) and corresponding fluorescent images (middle), showing changes in the intracellular distribution of LBs within a single cell before and after flow stimulation at 60 ␮l/min. A significant transport of LBs toward the cellular periphery is qualitatively observed from the fluorescent images captured at t ⫽ 0 and t ⫽ 20 min. Binary images (bottom) were extracted from the fluorescent images at an appropriate threshold value and then subtracted to obtain a new binary image representative of the relative changes (I0–I20) in LB locations (bottom, right).

diately after fusion of LBs with the PM, in combination with the intracellular distribution of LBs. For this purpose, we used a styryl dye, MM 1– 43, that binds to lipid LB content following formation of a fusion pore; note that tubule-like floating bodies resembling “tubular myelin” (TM) and protruding from exocytosed LBs stained with MM 1– 43 were observed at times in the bath medium. Such TMs consist of LB membranes that are often unpacked to form an organized network of crossing bilayers, where the TM-surfactant membranes then reach and

adsorb at the air-liquid interface to form films that fold upon compression to stabilize the airway lumen during cyclic compression-expansion dynamics (8). Compared with static conditions, we found under flow stimulation a significant increase in the average number of fusion events of LBs with the PM per cell for the MLE-12 population (see Fig. 4A, i). In contrast, our experiments reveal for A549 cells that this number remained statistically unaffected relative to static conditions for shear stress values ranging up to ⬃8 dyn/cm2 (see Fig. 4A, ii).

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L676

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

no flow

20 L/min

40 L/min

t=0

A

after 40 min

10 m

B (i)

(ii)

100 m

Fig. 3. A: morphological changes in cells subjected to variable flow rates. Note that cells subjected to a flow rate of 40 ␮l/min (or more) for 40 min resulted in significant alterations in morphologies, including shrinkage and round and oval-shaped appearances (highlighted in dashed red lines). Results for the quantitative evaluation of morphological changes in cell shape using cross-correlation techniques are elaborated in RESULTS. B: effects of fluid shear stress on cell viability. MLE-12 (i) and A549 (ii) cells subjected to a maximal flow rate of 100 ␮l/min for 20 min were tested for viability using fluorescent dyes, calcein acetoxymethyl (AM) in green and propidium iodide in red (see arrows).

Beyond this experimental WSS threshold value, the average number of LB fusion events per cell was found to be significantly lower for the two highest WSS magnitudes investigated (see Fig. 4A, ii). Compared with the trend seen for LB fusion events, we observed a positive correlation between flow rates applied and the relative incidence of intracellular shifts in the spatial distribution of LBs in A549 cells over the measured time interval (Fig. 4B); note again that, here, we quantify solely the incidence of such intracellular LB redistribution rather than the

direction of translocation per se (e.g., toward the periphery or the nucleus). Namely, we observe even in cells stimulated at the lowest experimental flow rate (i.e., 20 ␮l/min) that a significant amount of LBs translocate (Fig. 4B), without direct knowledge of direction. Qualitatively, we infer the likelihood of LB transport in the forward direction (i.e., toward the cellular periphery) as observed from fluorescent images (Fig. 2B, middle), where the intracellular redistribution of LBs in a single cell is shown at t ⫽ 0 and t ⫽ 20 min, respectively; careful examination of the image captured at t ⫽ 20 min

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

B

number of fusion events per cell

(ii)

2.5

2.5

relative incidence of LB resdistribution (%)

A (i)

number of fusion events per cell

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

*

2 1.5

**

** **

1 0.5 0

2 1.5

**

1

**

0.5 0

**

60 50

**

**

**

40

60

40 30 20 10 0

0

20

100

flow condition ( L/min)

Fig. 4. Effects of liquid flow-induced shear stress on the average number of LB fusion events (A) per cell (coinciding with exocytosed LBs) in MLE-12 (i) and A549 (ii) cells, after 20 min of flow stimulation, and on the relative incidence of LB redistribution (B) in A549 cells over time. Each value represents the mean of 5 different experiments; a total of over 250 cells were observed for each experimental condition to estimate the average number of LB fusion events per cell and the relative incidence (in percentage) of LB redistribution, respectively. Asterisks indicate statistically significant difference from the control; *P ⬍ 0.05, **P ⬍ 0.01.

L677

WSS can serve as a stimulatory factor for triggering the intracellular transport of LBs toward the periphery of A549 cells, it cannot promote or increase the fusion of LBs with the PM in such cells. Rather, it may instead potentially impair in vitro surfactant secretion above a given WSS threshold. Effects of shear stress on F-actin cytoskeleton. F-actin cytoskeleton is known to play a crucial role in the translocation of LBs and their exocytosis (36, 37, 45). Here, A549 cells treated with variable flow rates for 20 min were investigated qualitatively for the distribution pattern of actin filaments upon flow stimulation. Similar to static conditions, cells stimulated with 20 ␮l/min flow conditions showed organized and welldistributed filaments, in particular, around the periphery of the cell (Fig. 5). In contrast, cells subjected to flow rates of 40 ␮l/min and above exhibited fragmented, disorganized, and knot-like appearances of actin filaments in the cell cortex region (Fig. 5). Our qualitative assessment of actin filaments suggests that flow-induced shear stresses that range near or above the observed threshold values (i.e., 40 ␮l/min, corresponding to ⬃8 dyn/cm2) can lead to an F-actin disorganization. These results indicate that disorganization of actin filaments in A549 cells might be acting as a limiting factor for LB exocytosis above the experimental WSS threshold (Fig. 4A, ii). Influence of shear stress on intracellular calcium levels. Current knowledge suggests that calcium is a key regulator of exocytosis (5, 56). Hence, A549 cells were further examined to reveal the changes in the dynamic profile of [Ca2⫹]c upon flow stimulation. For the continuous monitoring of [Ca2⫹]c, cells loaded with a fluorescent indicator, Quest Fluo-8, were subjected to variable flow conditions and imaged to monitor changes in fluorescent intensity. Quest Fluo-8 AM ester is a membrane-permeable calcium indicator that rapidly gets hydrolyzed by cellular esterases inside live cells, yielding bright fluorescence intensity. Fig. 6A illustrates [Ca2⫹]c imaging in A549 cells stimulated with flow rates, 0, 20, 40, and 60 ␮l/min, respectively; the corresponding (mean) intensity profiles for a single cell representing each flow condition are plotted in Fig. 6B. Cells stimulated with 0 ␮l/min (static) and 20 ␮l/min flow conditions show a significant decay in fluorescence intensity over time, whereas cells subjected to flow rates of 40 and 60 ␮l/min display a visible rise in intensity within 1 min of flow stimulation, followed by a return to their baseline intensity value within 3 min of stimulation. Note that, before the initiation of WSS stimulation, a decay in fluorescence intensity profiles was briefly observed (⬃1 min) for all flow rates tested that may be attributed to photo-bleaching effects and was accounted for during quantitative measurements. Hence, our calcium imaging assay in A549 cells suggests that, below the experimental threshold WSS magnitude, flow-induced shear stress does not foreseeably induce an elevation of [Ca2⫹]c, whereas it can lead to well-defined peaks of [Ca2⫹]c for WSS values ranging at and above the experimental WSS threshold (⬃8 dyn/cm2). DISCUSSION

suggests the translocation of a significant number of LBs from the perinuclear region toward the cellular periphery. Taken together, our results suggest that flow-induced shear stress may act as a potential physical stimulus for LB fusion events (i.e., surfactant secretion) in MLE-12 cells. In contrast, although

The human lungs are adapted to facilitate gaseous exchange throughout postnatal life at the level of pulmonary alveoli (43). In particular, air-filled lungs are prone to form liquid plugs as a result of physiological disorders or therapeutic intervention (7, 48), where injury and death of ATII cells caused by fluid

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L678

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

20 L/min

no flow

40 L/min

10 m

60 L/min

100 L/min

Fig. 5. Fluorescent images showing the intracellular distribution of F-actin cytoskeleton. Here, A549 cells stimulated with variable flow rates were observed for changes in the distribution and organization of F-actin cytoskeleton using rhodamine-phalloidin.

WSS on underlying epithelial cells have been examined (7, 16, 20, 25, 48). However, the influence of WSS on surfactant secretion by ATII cells has not been investigated. Here, we have attempted to uncover the role of liquid flow-induced shear stresses on the mechanisms regulating surfactant secretion in ATII cells. Note that, unlike liquid plugs where large values of WSS arise among others due to sharp gradients in pressure at the liquid-gas interface between the propagating front of the plug and the neighboring air phase, here we have limited our study to fully immersed conditions in an effort to gain insight on the underlying mechanism of WSS on surfactant secretion. Although beyond the scope of the present study, only a coupling of mechanical stretching with fluid flow at an ALI would mimic more closely the true physiological process; until the present, such in vitro setups have been limited to studies of cell viability and detachment only (16). Moreover, because our microfluidic model is limited to investigations under fully immersed flow conditions only, the role of surface tension alterations that may arise at a fluid-fluid interface (i.e., airliquid) cannot be directly established. Because no single stable cell line expresses complete characteristics of in vivo ATII cells, two different models of ATII cells, i.e., MLE-12 and A549, were cultured into a monolayer inside microfluidic devices so as to mimic the alveolar epithelial cell lining at a physiological scale (16, 25). MLE-12 cells

show morphological characteristics and gene expression patterns similar to those seen in nonciliated bronchiolar and ATII cells (55). Also, MLE-12 cells express SP-C mRNA indicative of ATII cells in the adult mouse (55). However, such cell type does not share other characteristics with ATII cells, such as expression of SP-A mRNA or the presence of LBs (55). In contrast, the A549 cell line is probably the most frequently used alveolar epithelial model that shares some morphological and biochemical features of the human pulmonary ATII cells in situ (31). A549 cells contain multilamellar cytoplasmic inclusion bodies, like those typically found in human pulmonary ATII cells, and also produce surfactant phospholipids (34, 50, 51). However, expression of any of the surfactant proteins by A549 cells is still debated (34, 51). However, it has been shown that A549 cells exposed to air secrete surfactant into the liquid lining layer covering the cells, where a phospholipid film developed at the ALI; namely, this surfactant film may lower the surface tension of the liquid lining layer to values (28 mN/m) very close to those found in vivo (11). Here, cells were initially examined upon flow stimulation for the relative changes in the number of fusion events of LBs with the PM because the phenomenon is considered a key step of surfactant exocytosis. MLE-12 cells showed an increase in the number of LB fusion events compared with static conditions, in response to increasing WSS (Fig. 4A, i). In contrast, our

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L679

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

B t=0

1.06 min

2.17 min

5 min

50 m

(ii)

t=0

flow at 1.06 min

2.17 min

5 min

flow (20 l/min) flow (40 l/min) flow (60 l/min) static

35

fluorescence intensity (au)

A (i)

30 25 20 15 10 5 0

0

1

2

3

4

5

6

time (min)

(iii)

t=0

flow at 1.06 min

2.17 min

5 min

(iv)

t=0

flow at 1.06 min

2.17 min

5 min

Fig. 6. A: fluorescent imaging of cytosolic calcium ([Ca2⫹]c) levels in A549 cells subjected to variable flow rate conditions, 0 (static) (i), 20 (ii), 40 (iii), and 60 (iv) ␮l/min, are shown at different time instances. B: corresponding curves show changes in the level of [Ca2⫹]c (mean fluorescent intensity) during flow stimulation in a single cell for each flow condition. 3 independent experiments comprising triplicates of each condition were conducted to obtain the results.

measurements with A549 cells indicate that flow-mediated WSS values ranging up to ⬃8 dyn/cm2 have no foreseeable influence on the number of fusion events of LBs with the PM (Fig. 4A, ii), compared with static conditions, thereby suggesting that the total number of LBs involved in the detected fusion events might be similar under all conditions investigated. Beyond the experimental WSS threshold (i.e., ⬃8 dyn/cm2), we observed a marked decrease in LB exocytosis for shear stress values ranging between ⬃12 and 20 dyn/cm2. These results suggest that, for A549 cells, the step involving LB fusion with the PM is affected by a limiting factor associated with fluid shear stresses. We hypothesize a number of possible explanations to help interpret the contrasting behavior observed between MLE-12 (Fig. 4A, i) and A549 (Fig. 4A, ii) cell lines for the average number per cell of LB fusion events with the PM under increasing WSS stimulation. On the one hand, interspecies variations (i.e., murine vs. human cell lines) may lead to discrepancies in cellular responses stemming from distinct WSS experienced by the alveolar epithelium in vivo. Namely, WSS magnitudes are directly influenced by both morphological parameters (e.g., airway anatomy and size) and physiological flow parameters (e.g., tidal volume and breathing frequency); these are known to vary widely across species. Alternatively, intrinsic differences in surfactant protein expression between MLE-12 and A549 cell lines may be at the origin

of the discrepancies observed; whereas A549 cells are known to express SP-C (10, 34, 50, 51), MLEs in contrast express SP-A, SP-B, and SP-C (24, 55). In particular, some surfactant proteins can inhibit phospholipid secretion as for example with SP-A and phosphatidylcholine secretion (56). Finally, we note that MLE-12 cells have been shown to exhibit opposite responses to A549 cells when considering surfactant secretion patterns subject to in vitro cyclic stretching (i.e., mechanical strain) conditions (4). Our results for increasing WSS could potentially correspond to an analogous phenomenon under fluid shear stress stimulation. Next, molecular machineries required for the regulated secretion of surfactants in A549 cells were further explored, where A549 cells were examined for the relative occurrence of intracellular LB redistribution upon flow stimulation over a specific time interval (i.e., 20 min). Two distinct pools of secretory vesicles (e.g., LBs) have been identified in various cell types as the fundamental fusion machinery underlying exocytosis; these include a ”reserve pool,“ which represents most of the vesicle population, and a ”readily releasable pool⬙ most likely corresponding to vesicles already docked at the PM (33). Although several studies have suggested that in vitro primary ATII cells do not exhibit readily releasable pool of LBs (14, 17, 23), a recent study has revealed the presence of a limited pool of readily releasable vesicles in A549 cells (1).

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L680

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

Here, we have found that, in A549 cells, liquid flow stimulation triggers changes in the intracellular position of LBs belonging mostly to the reserve pool in a dose-dependent manner. In addition, our visual inspections of single cells further indicate that intracellular LB transport as a result of flow stimulation most likely occurs in the forward direction (Fig. 2B), suggesting that flow-induced shear stresses might serve as a physical stimulus for A549 cells to enhance the rate of transport of LBs toward the cellular periphery; this latter phenomenon is well established as one of the essential events underlying the secretory process (14, 15, 27, 52). In contrast, there was no corresponding rise observed in the number of fusion events in A549 cells for WSS values ranging up to ⬃8 dyn/cm2, thereby suggesting that the LBs involved in the detected fusion events probably belong to a ready-releasable pool, whereas the LBs transported from the reserve pool, as a result of the stimulus, did not take part in the fusion events. To gain further insight on the factor limiting LB fusion events in A549 cells, we analyzed the organization of F-actin filaments in ATII cells upon flow stimulation. The dual function of F-actin filaments in regulated secretory processes is well described, not only in other secretory cells such as neuron and endocrine cells (33), but also in alveolar type II cells (36, 40, 41, 49). Previous studies have suggested that the exocytosis process is accompanied by a fine reorganization (i.e., the local disassembly) of actin filaments that allows LBs from the reserve pool to gain access to the PM (41). Nevertheless, recent findings indicate that F-actin polymerization also plays a key role in a late postdocking step of exocytosis (36, 37). Our present results suggest that disorganization of F-actin filaments might act as a limiting factor for the LB fusion events in A549 cells (Fig. 5). Indeed, A549 cells under static conditions, as well as those stimulated for 20 min with a low WSS value of

⬃4 dyn/cm2, show polymerized and well-organized actin filaments, whereas the intracellular transport of LBs increases with growing WSS magnitudes, including beyond the WSS threshold. These contrasting observations suggest that those LBs transported from the reserve pool in A549 cells probably did not reach the docking sites at the PM due to the F-actin cytoskeleton acting as a physical barrier and may help explain the lack of any significant rise in the level of LB fusion events for flow stimulation below the threshold value. However, A549 cells subjected to WSS values ranging near or above the threshold exhibited disorganized and fragmented actin filaments, indicating that the concurrent rise of F-actin disorganization could be a limiting factor for no significant rise in the number of LB fusion events near the threshold. Instead, it appears that, for WSS above the experimental threshold, the organization of F-actin cytoskeleton was severely altered in association with a significant decrease in the number of LB fusion events in A549 cells. In a final step, we examined the level of [Ca2⫹]c in A549 cells under flow stimulation (Fig. 6). Ca2⫹ has long been recognized as one of the essential fusion machineries required for surfactant secretion (5, 6). Ca2⫹ plays a key role in regulating exocytosis processes as observed in various secretory cell types including ATII cells (13, 23, 47). Wirtz and Dobbs (56) were the first to show the time scale of calcium mobilization and exocytosis after a mechanical stretch of ATII cells. Furthermore, Frick et al. (18) found that strain-induced Ca2⫹ entry in ATII cells from the extracellular space is a prerequisite for Ca2⫹ release from intracellular stores (endoplasmic reticulum and LBs) and essential for LB fusion. Therefore, we examined changes in the level of [Ca2⫹]c upon flow stimulation in the presence of Ca2⫹-containing medium. For this purpose, cells were cultured to slightly lower levels of

Fig. 7. Quantifying morphological changes in cells with a normalized cross-correlation coefficient. Left: two phase-contrast images at time 0 and t ⫽ 20 min are extracted (example pair shown at 100 ␮l/min); middle: images are transformed into corresponding binary contours. Right: normalized correlation map is computed, from which the (maximum) correlation index value Rmax is extracted. AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

confluency than those used for other assays. Although it is likely that the nature or the extent of the response of cells against shear stress might vary at different levels of confluence, our objective here was to precisely analyze the intensity profile of a single cell. Namely, we found a significant decrease in the level of [Ca2⫹]c in A549 cells subjected to WSS values below the experimental threshold, indicating that this could be one of the possible reasons for no significant rise observed in the number of LB fusion events. In contrast, WSS values ranging near or above the threshold induced a high transient elevation of [Ca2⫹]c most likely due to the Ca2⫹ release from intracellular stores, suggesting that sufficient levels of [Ca2⫹]c were available in A549 cells to trigger the process of exocytosis. However, WSS magnitudes near or above the threshold led to no change or, in fact a significant decrease, in the number of LB fusion events, thus supporting the possibility of F-actin cytoskeleton acting at higher WSS values as a limiting factor for LB exocytosis in such cells. In summary, our in vitro assays suggest that ATII cells respond to liquid flow-induced shear stress in a differential manner depending on the cell type. Whereas MLE 12 cells exhibit a significant enhancement in the number of LB fusion events that are considered the ultimate step of surfactant exocytosis, A549 cells react instead by impairing LB exocytosis (i.e., fusion events) although they appear to propel their LBs toward their cell surface in response to flow stimulation. In analogy to the mechanical stretching-mediated surfactant secretion in ATII cells, shear stress-regulated secretory process is noted to be governed by the F-actin cytoskeleton and intracellular calcium concentration in A549 cells. Altogether, our findings suggest that liquid flow-induced shear stresses may play a nonnegligible role in regulating the molecular machinery required for surfactant secretion in ATII cells. However, we anticipate this effect to remain most likely minor relative to the primary stimulus of mechanical stretching. APPENDIX A: MORPHOLOGICAL CHANGES IN CELLS OVER TIME

To assess spatial changes in cell morphology over time in a quantitative matter, we have made use of a standard discrete crosscorrelation technique between two digital images (19), implemented in MatLab. This well-established technique enables one to quantify how closely two images resemble each other, by estimating a normalized correlation coefficient (or index) that compares the (maximum) correlation coefficient between an image at time 0 (ROIt⫽0) and t ⫽ 20 or 40 min (ROIt⫽20/40), relative to the corresponding (maximum) correlation coefficient obtained from an autocorrelation (i.e., crosscorrelation between ROIt⫽0 and itself). To assess how a given cell changes morphology over time, we first extract an image pair of a single cell at time 0 (ROIt⫽0) and t ⫽ 20 or 40 min (ROIt⫽20/40), from the original phase-contrast images (for example shown at 0 and 20 min in Fig. 7, left). Upon manual labeling, this image pair is then digitally transformed into a binary black and white image outlining the shape (or perimeter) of the cell (Fig. 7, middle). The two-dimensional discrete cross-correlation between images ROIt⫽0 and ROIt⫽20 of dimensions (M,N) is given by the expression: C(i, j) ⫽

L681

relation index (positioned at the specific location imax and jmax, and representing a shift in the position of the cell). Similarly, the autocorrelation function is defined as: Cauto(i, j) ⫽

M⫺1 N⫺1

兺 兺 ROIt⫽0(m, n) · ROIt⫽0(m ⫹ i, n ⫹ j),

m⫽0 n⫽0

from which we can extract the corresponding maximum correlation coefficient, max[Cauto(i,j)]. Finally, to compare quantitatively morphological changes occurring between time 0 and t ⫽ 20 (or 40) min, we introduce a normalized cross-correlation index R(i,j) ⫽ C(i,j)/max[Cauto(i,j)]; an example of the resulting image map R(i,j) is shown for an image pair in Fig. 7 (right). Recall that to produce a normalization index that is invariant to the segmentation size of the template, corresponding physically to changes in the size of a given cell between time 0 and 20/40, we select in practice the largest segmented template to construct the normalization index. In other words, the normalization template is chosen at time 0, unless it is larger at t ⫽ 20/40 (i.e., the 2D image of the cell covers a larger area at t ⫽ 20/40 compared with time 0). This may be formally written as max[Cauto(i,j)] ⫽ max(max[Cauto(i,j)t⫽0]; max[Cauto(i,j) t⫽20/40]). The index R(i,j) introduced here is slightly different from the traditional normalized cross-correlation definition (19), where the cross-correlation operation is done at every step (i.e., pixel) by subtracting the image mean and dividing by the image standard deviation. Because we are only interested in the best-matched normalized correlation index (rather than locating the exact shift in the spatial position of the cell), we can extract the quantity Rmax ⫽ max[R(i,j)]. Note that a hypothetical value of Rmax ⫽ 1 would indicate that an image at t ⫽ 20/40 min is perfectly identical to that at t ⫽ 0. In practice, values of Rmax ⬎ 0.9 indicate that two images are highly similar. ACKNOWLEDGMENTS The authors thank Dr. Limor Minai (Technion, Israel) for helpful discussions on cell biology and Dr. Raphael Sznitman (EPFL, Switzerland) for fruitful discussions on image processing. Microfabrication was conducted at the Micro-Nano fabrication Unit (Technion). Present address for S. Mahto: Department of Electrical and Computer Engineering, University of Calgary, Calgary, Alberta, Canada. GRANTS This work was supported in part by a Technion postdoctoral fellowship (Dr. Sanjeev K. Mahto), the European Commission (FP7 Program) through a Career Integration Grant (PCIG0-GA-2011-293604), as well as by the Israel Science Foundation (ISF grant nr. 990/12), Research Grant Award No. PGA 1302 from the Environment and Health Fund, Israel. The authors acknowledge support from the Nevet grant program of the Russel Berrie Nanotechnology Institute and the Technion Center of Excellence in Exposure and Environmental Health (Technion). DISCLOSURES No conflicts of interest, financial or otherwise, are declared by the authors. AUTHOR CONTRIBUTIONS Author contributions: S.K.M. and J.S. conception and design of research; S.K.M. performed experiments; S.K.M., A.G., and J.S. analyzed data; S.K.M., J.T.-K., and J.S. interpreted results of experiments; S.K.M. prepared figures; S.K.M. and J.S. drafted manuscript; S.K.M., J.T.-K., B.R.-R., and J.S. edited and revised manuscript; S.K.M. and J.S. approved final version of manuscript.

M⫺1 N⫺1

兺 兺 ROIt⫽0(m, n) · ROIt⫽20(m ⫹ i, n ⫹ j),

m⫽0 n⫽0

where C(i,j) is the correlation coefficient at any pixel location i and j, with 0 ⱕ i ⱕ M and 0 ⱕ j ⱕ N. The maximum correlation coefficient, max[C(i,j)], denotes the quantitative value for the best-matched cor-

REFERENCES 1. Akopova I, Tatur S, Grygorczyk M, Luchowski R, Gryczynski I, Gryczynski Z, Borejdo J, Grygorczyk R. Imaging exocytosis of ATPcontaining vesicles with TIRF microscopy in lung epithelial A549 cells. Purinergic Signal 8: 59 –70, 2011.

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

L682

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION

2. Andreeva AV, Kutuzov MA, Voyno-Yasenetskaya TA. Regulation of surfactant secretion in alveolar type II cells. Am J Physiol Lung Cell Mol Physiol 293: L259 –L271, 2007. 3. Arold SP, Bartolak-Suki E, Suki B. Variable stretch pattern enhances surfactant secretion in alveolar type II cells in culture. Am J Physiol Lung Cell Mol Physiol 296: L574 –L581, 2009. 4. Arold SP, Wong J, Suki B. Design of a new stretching apparatus and the effects of cyclic strain and substratum on mouse lung epithelial-12 cells. Ann Biomed Eng 35: 1156 –1164, 2007. 5. Ashino Y, Ying X, Dobbs LG, Bhattacharya J. [Ca2⫹]i oscillations regulate type II cell exocytosis in the pulmonary alveolus. Am J Physiol Lung Cell Mol Physiol 279: L5–L13, 2000. 6. Barclay JW, Morgan A, Burgoyne RD. Calcium-dependent regulation of exocytosis. Cell Calcium 38: 343–353, 2005. 7. Bilek AM, Dee KC, Gaver DP. Mechanisms of surface-tension-induced epithelial cell damage in a model of pulmonary airway reopening. J Appl Physiol 94: 770 –783, 2003. 8. Blanco O, Pérez-Gil J. Biochemical and pharmacological differences between preparations of exogenous natural surfactant used to treat Respiratory Distress Syndrome: role of the different components in an efficient pulmonary surfactant. Eur J Pharmacol 568: 1–15, 2007. 9. Bland RD, Nielson DW. Developmental changes in lung epithelial ion transport and liquid movement. Annu Rev Physiol 54: 373–394, 1992. 10. Blank F, Rothen-Rutishauser B, Schurch S, Gehr P. An optimized in vitro model of the respiratory tract wall to study particle cell interactions. J Aerosol Med 19: 392–405, 2006. 11. Bruus H. Theoretical Microfluidics. Oxford, UK: Oxford University, 2007. 12. Chander A, Fisher AB. Regulation of lung surfactant secretion. Am J Physiol Lung Cell Mol Physiol 258: L241–L253, 1990. 13. Chow RH, Klingauf J, Neher E. Time course of Ca2⫹ concentration triggering exocytosis in neuroendocrine cells. Proc Natl Acad Sci USA 91: 12765–12769, 1994. 14. Dietl P, Haller T, Mair N, Frick M. Mechanisms of surfactant exocytosis in alveolar type II cells in vitro and in vivo. Physiology 16: 239 –243, 2001. 15. Dietl P, Liss B, Felder E, Miklavc P, Wirtz H. Lamellar body exocytosis by cell stretch or purinergic stimulation: possible physiological roles, messengers and mechanisms. Cell Physiol Biochem 25: 1–12, 2010. 16. Douville NJ, Zamankhan P, Tung YC, Li R, Vaughan BL, Tai CF, White J, Christensen PJ, Grotberg JB, Takayama S. Combination of fluid and solid mechanical stresses contribute to cell death and detachment in a microfluidic alveolar model. Lab Chip 11: 609, 2011. 17. Frick M, Eschertzhuber S, Haller T, Mair N, Dietl P. Secretion in alveolar type II cells at the interface of constitutive and regulated exocytosis. Am J Respir Cell Mol Biol 25: 306 –315, 2001. 18. Frick M, Bertocchi C, Jennings P, Haller T, Mair N, Singer W, Pfaller W, Ritsch-Marte M, Dietl P. Ca2⫹ entry is essential for cell straininduced lamellar body fusion in isolated rat type II pneumocytes. Am J Physiol Lung Cell Mol Physiol 286: L210 –L220, 2004. 19. Gonzalez RC. Digital Image Processing Using MATLAB. 2nd ed. New York, NY: Gatesmark, 2009. 20. Grotberg JB. Respiratory fluid mechanics. Phys Fluids 23: 021301021301-15, 2011. 21. Haefeli-Bleuer B, Weibel ER. Morphometry of the human pulmonary acinus. Anat Rec 220: 401–414, 1988. 22. Haller T, Dietl P, Pfaller K, Frick M, Mair N, Paulmichl M, Hess MW, Fürst J, Maly K. Fusion pore expansion is a slow, discontinuous, and Ca2⫹-dependent process regulating secretion from alveolar type II cells. J Cell Biol 155: 279 –290, 2001. 23. Haller T, Ortmayr J, Friedrich F, Völkl H, Dietl P. Dynamics of surfactant release in alveolar type IIcells. Proc Natl Acad Sci USA 95: 1579 –1584, 1998. 24. Horowitiz AD, Moussavian B, Whitsett JA. Roles of SP-A, SP-B, and SP-C in modulation of lipid uptake by pulmonary epithelial cells in vitro. Am J Physiol Lung Cell Mol Physiol 270: L69 –L79, 1996. 25. Huh D, Fujioka H, Tung YC, Futai N, Paine R 3rd, Grotberg JB, Takayama S. Acoustically detectable cellular-level lung injury induced by fluid mechanical stresses in microfluidic airway systems. Proc Natl Acad Sci USA 104: 18886 –18891, 2007. 26. Huh D, Hamilton GA, Ingber DE. From 3D cell culture to organs-onchips. Trends Cell Biol 21: 745–754, 2011. 27. Johnson NF. Release of lamellar bodies from alveolar type 2 cells. Thorax 35: 192–197, 1980.

28. Kolhar P, Anselmo AC, Gupta V, Pant K, Prabhakarpandian B, Ruoslahti E, Mitragotri S. Using shape effects to target antibody-coated nanoparticles to lung and brain endothelium. Proc Natl Acad Sci USA 110: 10753–10758, 2013. 29. Lieber M, Smith B, Szakal A, Nelson-Rees W, Todaro G. A continuous tumor-cell line from a human lung carcinoma with properties of type II alveolar epithelial cells. Int J Cancer 17: 62–70, 1976. 30. Liu M, Tanswell AK, Post M. Mechanical force-induced signal transduction in lung cells. Am J Physiol Lung Cell Mol Physiol 277: L667– L683, 1999. 31. Mahto SK, Tenenbaum-Katan J, Sznitman J. Respiratory physiology on a chip. Scientifica 2012: 364054, 2012. 32. Mahto SK, Yoon TH, Shin H, Rhee SW. Multicompartmented microfluidic device for characterization of dose-dependent cadmium cytotoxicity in BALB/3T3 fibroblast cells. Biomed Microdevices 11: 401–411, 2009. 33. Malacombe M, Bader MF, Gasman S. Exocytosis in neuroendocrine cells: new tasks for actin. Biochim Biophys Acta 1763: 1175–1183, 2006. 34. Mason RJ, Williams MC. Phospholipid composition and ultrastructure of A549 cells and other cultured pulmonary epithelial cells of presumed type II cell origin. Biochim Biophys Acta 617: 36 –50, 1980. 35. McDonald JC, Duffy DC, Anderson JR, Chiu DT, Wu H, Schueller OJ, Whitesides GM. Fabrication of microfluidic systems in poly(dimethylsiloxane). Electrophoresis 21: 27–40, 2000. 36. Miklavc P, Hecht E, Hobi N, Wittekindt OH, Dietl P, Kranz C, Frick M. Actin coating and compression of fused secretory vesicles are essential for surfactant secretion—a role for Rho, formins and myosin II. J Cell Sci 125: 2765–2774, 2012. 37. Miklavc P, Wittekindt OH, Felder E, Dietl P. Ca2⫹-dependent actin coating of lamellar bodies after exocytotic fusion: a prerequisite for content release or kiss-and-run. Ann NY Acad Sci 1152: 43–52, 2009. 38. Ochs M, Weibel ER. Functional design of the human lung for gas exchange. In Fishman’s Pulmonary Diseases And Disorders, edited by AP Fishman, JA Elias, JA Fishman. New York, NY: McGraw Hill, 2007, pp. 23– 69. 39. Ohashi T, Sato M. Remodeling of vascular endothelial cells exposed to fluid shear stress: experimental and numerical approach. Fluid Dyn Res 37: 40 –59, 2005. 40. Porat-Shliom N, Milberg O, Masedunskas A, Weigert R. Multiple roles for the actin cytoskeleton during regulated exocytosis. Cell Mol Life Sci 70: 2099 –2121, 2013. 41. Rose F, Kürth-Landwehr C, Sibelius ULF, Reuner KH, Aktories K, Seeger W, Grimminger F. Role of actin depolymerization in the surfactant secretory response of alveolar epithelial type II cells. Am J Respir Crit Care Med 159: 206 –212, 1999. 42. Rothen-Rutishauser B, Jud C, Wick P, Clift MJ, Fink A. Human epithelial cells in vitro—Are they an advantageous tool to help understand the nanomaterial-biological barrier interaction? Euro Nanotox Lett 4: 1–20, 2012. 43. Ryan US, Ryan JW, Smith DS. Alveolar type II cells: studies on the mode of release of lamellar bodies. Tissue Cell 7: 587–599, 1975. 44. Scott JE, Yang SY, Stanik E, Anderson JE. Influence of strain on [3H]thymidine incorporation, surfactant-related phospholipids synthesis, and cAMP levels in fetal type II alveolar cells. Am J Respir Cell Mol Biol 8: 258 –265, 1993. 45. Singh TK, Abonyo B, Narasaraju TA, Liu L. Reorganization of cytoskeleton during surfactant secretion in lung type II cells: a role of annexin II. Cell Signal 16: 63–70, 2004. 46. Stork CJ, Li YV. Measuring cell viability with membrane impermeable zinc fluorescent indicator. J Neurosci Methods 155: 180 –186, 2006. 47. Südhof TC. The synaptic vesicle cycle: a cascade of protein-protein interactions. Nature 375: 645–653, 1995. 48. Tavana H, Kuo CH, Lee QY, Mosadegh B, Huh D, Christensen PJ, Grotberg JB, Takayama S. Dynamics of liquid plugs of buffer and surfactant solutions in a micro-engineered pulmonary airway model. Langmuir 26: 3744 –3752, 2010. 49. Tsilibary EC, Williams MC. Actin and secretion of surfactant. J Histochem Cytochem 31: 1298 –1304, 1983. 50. Wang D, Haviland DL, Burns AR, Zsigmond E, Wetsel RA. A pure population of lung alveolar epithelial type II cells derived from human embryonic stem cells. Proc Natl Acad Sci USA 104: 4449 –4454, 2007. 51. Wang J, Edeen K, Manzer R, Chang Y, Wang S, Chen X, Funk CJ, Cosgrove GP, Fang X, Mason RJ. Differentiated human alveolar epi-

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

MICROFLUIDIC SHEAR STRESS-REGULATED SURFACTANT SECRETION thelial cells and reversibility of their phenotype in vitro. Am J Respir Cell Mol Biol 36: 661–668, 2007. 52. Weaver TE, Na CL, Stahlman M. Biogenesis of lamellar bodies, lysosome-related organelles involved in storage and secretion of pulmonary surfactant. Semin Cell Dev Biol 13: 263–270, 2002. 53. Weibel ER, Gomez DM. Architecture of the human lung use of quantitative methods establishes fundamental relations between size and number of lung structures. Science 137: 577–585, 1962. 54. Weibel ER. Morphometry of the Human Lung. New York: Academic, 1963.

L683

55. Wikenheiser KA, Vorbroker DK, Rice WR, Clark JC, Bachurski CJ, Oie HK, Whitsett JA. Production of immortalized distal respiratory epithelial cell lines from surfactant protein C/simian virus 40 large tumor antigen transgenic mice. Proc Natl Acad Sci USA 90: 11029 – 11033, 1993. 56. Wirtz HR, Dobbs LG. Calcium mobilization and exocytosis after one mechanical stretch of lung epithelial cells. Science 250: 1266 –1269, 1990. 57. Yalcin HC, Perry SF, Ghadiali SN. Influence of airway diameter and cell confluence on epithelial cell injury in an in vitro model of airway reopening. J Appl Physiol 103: 1796 –1807, 2007.

AJP-Lung Cell Mol Physiol • doi:10.1152/ajplung.00106.2013 • www.ajplung.org

Copyright of American Journal of Physiology: Lung Cellular & Molecular Physiology is the property of American Physiological Society and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.