www.sciencemag.org/cgi/content/full/328/5986/1662/DC1

Supporting Online Material for

Reconstituting Organ-Level Lung Functions on a Chip

Dongeun Huh, Benjamin D. Matthews, Akiko Mammoto, Martín Montoya-Zavala, Hong Yuan Hsin, Donald E. Ingber*

*To whom correspondence should be addressed. E-mail: don.ingber@wyss.harvard.edu

Published 25 June 2010, Science 328, 1662 (2010) DOI: 10.1126/science.1188302

This PDF file includes:

Materials and Methods SOM Text Figs. S1 to S11 Table S1 References

Other Supporting Online Material for this manuscript includes the following: (available at www.sciencemag.org/cgi/content/full/328/5986/1662/DC1)

Movies S1 to S9

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Materials and Methods Device fabrication. The upper and lower layers of the microfluidic device shown in Fig. 1D were produced by casting PDMS prepolymer against a photolithographically prepared master that contains a positive relief of parallel microchannels made of photoresist (SU8-50, MicroChem). The weight ratio of PDMS base to curing agent was 15:1. The cross-sectional size of the microchannels is 400 μm (width) X 70 μm (height) for the central culture channels and 200 μm (width) X 70 μm (height) for the side channels. Thin

microporous PDMS membranes were generated by spin-coating PDMS prepolymer (15:1) on a silanized wafer that has an array of 50 μm-tall pentagonal posts fabricated by using standard photolithographic techniques. Spin-coating at 2500 rpm for 10 minutes produced 10 μm-thick PDMS membranes with pentagonal through-holes. After curing at 65ºC overnight, the membrane surface was briefly treated with corona plasma generated by a hand-held corona treater (Electro-Technic Products) and brought in conformal contact with the upper PDMS substrate to achieve irreversible bonding between the layers. After overnight incubation at 65ºC, the bottom surface of the membrane was treated with corona and permanently bonded to the lower PDMS layer after careful manual alignment. For PDMS etching, tetrabutylammonium fluoride (TBAF) was mixed with N-methylpyrrolidinone (NMP) at a volumetric ratio of 1:3 (TBAF: NMP) and used as an etchant (S1). An etching solution was introduced into the side microchannels through inlet reservoirs by using hydrostatic pressure or vacuum suction at the outlet ports. The flow of etchant was driven at a constant flow rate of ~ 200 μl/min. The membrane layers containing pentagonal holes in the side channels

completely etched away within 2 minutes. Etching continued until the thickness of the

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PDMS walls between the side chambers from the central culture channels became thinner than 30 μm. After etching, the side chambers were washed thoroughly with NMP for at least 3 minutes to remove any remaining PDMS etchant. Microfluidic cell culture. Human pulmonary microvascular endothelial cells (Clonetics) were cultured in EBM-2 medium supplemented with 5% FBS and growth factors according to the manufacturer’s protocols. Alveolar epithelial cells used in this work were NCI H441 (ATCC). A549 (ATCC), and E10 (a gift from Dr. Randall Ruch at the University of Toledo) were used to visualize cell stretching in our microfluidic device and to validate culture conditions during microfluidic cell culture. These cells were grown in RPMI-1640 (NCI H441), Ham’s F12K (A549), or CMRL-1066 (E10) medium, respectively, supplemented with 10% FBS, L-glutamine (0.292 mg/ml), penicillin (100 U/ml), and streptomycin (100 μg/ml). The cells were maintained at 37ºC in a humidified incubator under 5% CO2 in air. Prior to cell seeding, microfluidic devices were sterilized by UV irradiation, and the porous membranes embedded in the central culture channels were coated with collagen gel (3 μg/ml) or fibronectin (5 μg/ml in carbonate buffer). Alveolar epithelial cells were seeded into the upper channel at approximately 2X104 cells per cm2 and allowed to attach to the membrane surface for 2 hours under static conditions. The attached cells were then perfused with culture medium by a syringe pump at a volumetric flow rate of 20 μl/hr. After 24 hours, the flow of culture medium was stopped, and the microfluidic device was inverted to seed endothelial cells onto the opposite side of the membrane. Because the membrane surface was coated with collagen, the cells did not go through the membrane pores during cell seeding.

After cell attachment, steady flows of culture media were driven in both the upper and lower channels at 20 μl/hr (fluid shear stress = 0.1 dyne/cm2). The cells in both microchambers were grown to confluence within 5 days. Once alveolar epithelial cells reached confluence, they were treated with culture medium containing 1 μM of glucocorticoid dexamethasone (Sigma) to promote the formation of tight junctions (S2). For air-liquid interface culture, culture medium was gently aspirated from the upper channel on Day 5, and a 50:50 mixture of epithelial and endothelial medium was introduced into the lower channel to feed the alveolar epithelial cells on their basolateral side. The epithelial cells were grown at an air-liquid interface for 16 days. Microfluidic culture was maintained at 37ºC in a humidified incubator with 5% CO2 in air.

The alveolar-capillary unit structure was visualized by confocal imaging of a

monolayer of alveolar epithelial cells labeled with CellTracker Green CMFDA (Invitrogen) closely apposed to a layer of microvascular endothelial cells stained with CellTracker Red CMTPX (Invitrogen); representative images are shown in Fig. 2A (upper and middle insets). The cells were labeled fluorescently with these vital dyes before being introduced into the microchannels, and the images recorded using confocal microscopy after 3 weeks of culture were carefully examined to show that the cells remained in their original compartments without migrating across the membrane. Mechanical stretching. To visualize membrane stretching in our microfluidic system, we created arrays of microscopic fluorescent spots on PDMS membranes by depositing small volumes of fluorescent nanoparticle suspensions at pre-defined positions using an

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automated high-precision NanoeNablerTM printer (BioForce Nanosciences) (Fig. 2B and Movie S2). Prior to mechanical stretching, the outlet holes of the side chambers were blocked, and the inlet ports were connected to a computer-controlled vacuum pump via vacuum-resistant tubing. Cyclic stretching was achieved by applying vacuum to the two side chambers simultaneously in a cyclic fashion. To mimic physiological breathing motion, the alveolar-capillary barriers formed in the microfluidic device were stretched with 5 ~ 15% strain at a frequency of 0.2 Hz (sinusoidal waveform). To examine the effect of pre-conditioning with strain on barrier functions (fig. S1), cells were subjected to cyclic stretch for varying amounts of time that have been reported in the literature to be relevant for studies of lung cell function (S3-S7), prior to the measurement of barrier permeability. Cells were stretched for 5 hours and 5 days to investigate short-term and long-term priming effects, respectively. Immunostaining of junctional complexes. To stain for Occludin, a transmembrane protein localized to epithelial tight junctions, alveolar epithelial cells in the upper microchannel were washed with phosphate buffered saline (PBS), fixed with 4% paraformaldehyde (PFA) in PBS for 20 minutes, washed again with PBS, and permeabilized with 0.3% Triton X-100 in PBS for 5 minutes. After washing and blocking with 1% bovine serum albumin (BSA) in PBS for 1 hour, the cells were incubated with Alexa 594-conjugated mouse anti-human occludin antibody (Invitrogen) for 1 hour and washed with PBS. For staining of vascular endothelial (VE)-cadherin, microvascular endothelial cells in the microfluidic device were fixed, permeabilized, and blocked using the same methods described above. The cells were then incubated with mouse anti-human VE-cadherin antibody for 1 hour and washed with PBS. Subsequently, Alexa 594-conjugated secondary antibody was added and incubated for 1 hour before fluorescence imaging. Permeability assay. The permeability of the alveolar-capillary barrier was assessed by measuring the rate of fluorescein isothiocyanate (FITC)-conjugated albumin transport from the upper alveolar channel to the lower vascular compartment. 1 mg/mL of FITC-albumin in epithelial medium was injected into the upper channel, and albumin transport across the barrier was determined by serially sampling liquid flowing out of the lower channel due to a continuous flow driven at 30 μL/hr and measuring its fluorescence intensity. A standard curve was established using known concentrations of FITC-albumin solution to correlate the intensity of fluorescence with concentration. Trans-bilayer electrical resistance (TER) measurements. TER was measured using a volt-ohm meter modified with wire silver/silver-chloride (Ag/AgCl) electrodes. The two wire electrodes were inserted into the inlet of the upper alveolar compartment and the outlet of the lower vascular channel, respectively. The contribution of the porous membrane and culture medium was accounted for by subtracting the blank resistance of the membrane without cells. TER was evaluated every 3 days during air-liquid interface culture and used as a measure of barrier integrity. Cellular treatment with nanomaterials and ROS detection. To detect intracellular production and accumulation of ROS, alveolar epithelial cells and microvascular endothelial cells in our device were briefly washed with serum-free medium after air-liquid

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interface culture and incubated with the ROS indicator dye, 5-(and 6)-chloromethyl-2’,7’-dichlorodihydrofluorescein diacetate (CM-H2DCFDA) (Invitrogen) at 37ºC for 1 hour. CM-H2DCFDA was used at a final working concentration of 10 μM in serum-free culture medium (S8). After washing, nanomaterials suspended in lung cell culture medium were injected into the alveolar space. Subsequently, the solution in the alveolar microchannel was gently aspirated to form a thin liquid layer containing nanomaterials on the apical surface of the epithelial cells. The epithelial layer was exposed to different types of nanomaterials at desired concentrations and times. Silica nanoparticles, carboxylated quantum dots, and carboxylated superparamagnetic iron nanoparticles were obtained from Sigma (Ludox LS colloidal silica; diameter ~ 12 nm; negatively charged; specific gravity = 1.21), Invitrogen (Qdot 605 ITK carboxyl quantum dots; diameter ~ 16 nm), and Kisker Biotechnology (Kisker PMC-50; diameter ~ 50 nm), respectively. Polystyrene nanoparticles were purchased from Invitrogen (FluoSpheres Fluorescent Microspheres; carboxylate-modified microspheres with diameters of 500 nm, 200 nm, 100 nm, and 20 nm; amine-modified microspheres with a diameter of 200 nm). Single-walled carbon nanotubes (SWNTs) and gold nanotubes were a gift from Dr. Peter Cherukuri at the University of Texas M. D. Anderson Cancer Center (diameter and length of the SWNTs are 1 nm and 300 nm, respectively). The concentration of nanomaterials tested was determined based on the range of nanomaterial concentrations that were previously reported to produce physiologically significant effects in the literature. The microfluidic device was kept in a cell culture incubator during nanomaterial exposure. ROS generation (Figs. 4B and 4C) was quantified by the intensity of intracellular fluorescence averaged over 12 different observation areas within the microchannel from three separate experiments. Data shown in Figs. 4A and 4D were gathered from two separate experiments using air-liquid interface culture, and similar results were obtained in three additional experiments when nanoparticles were introduced in culture medium. Obtained data were represented as mean ± SEM, and their statistical significance was determined by analysis of variance (ANOVA) followed by post hoc Tukey’s multiple comparison test. Immunohistochemical detection of ICAM-1. After TNF-α stimulation of the alveolar epithelium, microvascular endothelial cells on the opposite side of the culture membrane were fixed with 4% PFA in PBS, permeabilized with 0.3% Triton X-100 in PBS, and blocked with 1% BSA in PBS. Subsequently, the cells were incubated with mouse anti-human ICAM-1 antibody (S9) for 1 hour, washed with PBS, and incubated with Alexa 594-conjugated secondary antibody for 1 hour for optical detection of fluorescently labeled ICAM-1 on the endothelial surface. Isolation of human neutrophils. Human neutrophils were isolated from sodium citrated whole blood drawn from healthy consenting donors by using the RosetteSep® human granulocyte enrichment kit (StemCell Technologies) according to the procedure provided by the manufacturer. Isolated neutrophils were labeled with Cell Tracker Red CMTPX (Invitrogen) and injected into the lower microchannel in medium at a volumetric flow rate of 50 ~ 100 μL/hr using a syringe pump. Bacterial transformation and microfluidic infection. E. coli XL1-Blue competent cells were transformed with pGLO plasmid (Bio-Rad) in the presence of ampicillin according to the manufacturer’s protocol (Stratagene). Arabinose was then used to induce the

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expression of the pGLO plasmid as demonstrated by the presence of GFP fluorescence. The transformed bacteria suspended in epithelial culture medium were introduced into the upper microchannel via a syringe pump and incubated for 5 hours to infect the alveolar epithelium. Animal Work All experimental animal protocols were approved by the Institutional Animal Care and Use Committee at Children’s Hospital Boston and Harvard Medical School. Anesthetized mice were euthanized after thoracotomy by exsanguination prior to pulmonary artery (PA) and left atrial (LA) cannulation.

Mouse tracheal nebulization catheter and system. A tracheal nebulization catheter system (see fig. S11) was constructed using a customized Aeroprobe© Intracorporeal Nebulizing Catheter (INC, Trudell Manufacturing International, London, Canada). The Aeroprobe© INC is secured within the nebulization catheter system by virtue of a Touhy Borst Luer Lock Adapter (Trudell Manufacturing International) which appears at the far left of fig. S10B. The catheter tip is positioned at the outlet of an endotracheal tube fashioned from a 22G stainless steel needle (Becton, Dickinson and Company, NJ) seen on the far right in Fig S10B. The endotracheal tube is attached proximally to a plastic Y-connector which allows simultaneous nebulization and mechanical ventilation (see ex vivo ventilation perfusion below). A LABneb© Catheter Control Unit (CCU, Trudell Manufacturing International) was used to deliver 5 x 5 ms pulses (at 100 psi using compressed air) of nebulized nanoparticles (FluoSpheres® carboxylate-modified microspheres, 20 nm, red fluorescent (580/605) , 2% solids suspended in 1 mL PBS, Invitrogen Life Science, CA) through the nebulization catheter and into the animal lungs. Pulses were timed manually to correspond with the inspiratory phase of consecutive ventilator breaths. Ex vivo lung ventilation perfusion experiments. 8 week-old male C57BL/6 mice (The Jackson Laboratory, Bar Harbor, ME) were weighed and then anesthetized with Avertin (200 mg/kg IP). Ex vivo ventilation and perfusion of the mouse lung was facilitated by the IL1 ex vivo mouse lung ventilation perfusion system (Harvard Apparatus, Natick, MA). The trachea was incised via surgical tracheotomy, and a specialized tracheal nebulization catheter system was secured inside the trachea. The lungs were subsequently ventilated at a rate of 60 breaths per minute, with a Peak Inspiratory Pressure (Pip) of 10 cm H2O and a Positive End Expiratory Pressure (Peep) of 3 cm H2O with compressed air using a mouse ventilator (VCM-R, Hugo Sachs Elektroniks, Germany). Following the initiation of mechanical ventilation, the chest was opened via thoracotomy, and heparin (100 IU) was injected into the right ventricle. After 30 seconds, the thoracic aorta and superior vena cava were cut and the animal ex-sanguinated. A suture was placed around the pulmonary artery and aorta. Cannulae (0.86 mm ID, 1.27 mm OD, Harvard Apparatus, Natick, MA)

were placed in the pulmonary artery (PA) and left atrium (LA), and lungs were perfused with RPMI-1640 with 4% Bovine Albumin (ProbuminTM Reagent Grade, Billerica, MA) and 0.7 g NaCl/500 mL68 via a roller pump (ISM 834C, Ismatec SA, Switzerland) set at a constant flow rate of 0.5 ml/min in a recirculating system with a system volume of 6 mL. For 10 minutes following initiation of perfusion, the lungs were observed for leakage and or

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obstruction to perfusion or airflow. If leakage or obstruction were observed, the experiment was aborted. If not observed, the initial 2 ml of perfusate, which contained residual blood cells and plasma, were discarded and not recirculated. A 200 μL sample of perfusate was then drawn from the LA cannula to serve as baseline for later quantification of nanoparticle content. 5 pulses of nebulized nanoparticles were then delivered to the mouse lung as described above. 200 μL perfusate volumes were sampled every 10 minutes from the LA catheter for later quantification of nanoparticle content until the end of the experiment at 60 minutes post nebulization. The lungs from the control groups were ventilated for the entire 60 minutes at the PIP and Peep as indicated above, whereas the ventilator was turned off in the non-ventilated groups immediately following nebulization. Perfusate and lung temperatures were maintained at 33°C by the IL1 system using a temperature regulated recirculating water bath. Pulmonary arterial and venous pressures and airway flow and pressures were recorded with dedicated Type 379 vascular pressure and DLP2.5 flow and MPX Type 399/2 airway pressure transducers and TAM-A amplifiers (Hugo Sachs Elektroniks, Germany). Vascular pressures were zeroed at the midlung level prior to each experiment and recorded using Polyview16© software (Grass Technologies, West Warwick, RI) running on a desktop PC running Windows XP SP2 (Microsoft Corporation, Redmond, WA). The nebulization and ex vivo ventilation perfusion experiments were performed inside a conventional laboratory fume hood. Lung weights and tissue section histology. At the end of each experiment, the lungs were fixed by vascular perfusion with 4% paraformaldehyde using a syringe pump (Braintree Scientific, Braintree, MA), dissected and weighed. Lungs were subsequently incubated overnight in 4% paraformaldehyde at 4 deg C, then washed in 70% ethanol, and paraffin embedded using standard techniques. Lungs were then cut into 5 μm thick sections and processed either for H&E staining using standard techniques, or stained with phalloidin (Alexa 488, Molecular Probes) to identify F-actin and then mounted with Prolong Gold anti fade mounting reagent with DAPI (Invitrogen) to stain for nuclei. Quantification of nanoparticle translocation into ex vivo perfusate system. To quantify nanoparticle (20 nm) translocation from the alveoli into the perfusate circulation, 200 μL samples of perfusate were collected from the LA catheter every 10 minutes for 60 minutes during the ex vivo experiments described above and following nebulization. 1 μL aliquots were taken from each sample and placed onto a glass slide (Corning Incorporated, Corning, NY) and allowed to dry. The dried aliquots were covered with a glass coverslip (No. 0 thickness, Corning Incorporated) and sealed with clear nail polish. The slides were analyzed for particle content on a Nikon Diaphot 300 fluorescence microscope (Nikon, Japan) using a CCD camera (Hamamatsu, Japan) controlled by IPLab software (version 3.2.4, Scanalytics, VA). 10 images were captured per slide at locations that were pre-determined by a macro written in IPLab. Total number of particles per μm2 was determined for each sample by dividing the total number of nanoparticles seen in the 10 images by the total area scanned (4.42 x 105 μm2).

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Supporting Text Effect of flow pulsatility. Cyclic stretch generates pulsatility in the fluid flow and a cyclic variation in fluid shear stress. To determine whether this effect is relevant, we used a simple scaling analysis based on the Womersley number, which is a dimensionless parameter used to describe unsteady pulsatile fluid flow. The Womersley number (α) is defined as α = h(ω/ν)1/2, where h is the channel height, ω is the frequency of pulsatile flow, and ν is the kinematic viscosity. For our microfluidic device, α is estimated to be very small (0.022) and therefore, the unsteady effects on shear stress are negligible.

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Fig. S1. (A) Cellular viability was maintained over 85% throughout the air-liquid interface culture period (16 days) in our microfluidic device (open rectangles, epithelium; closed rectangles, endothelium). Alveolar epithelial cells were exposed to air on Day 5 (data represent the mean ± SEM of three separate experiments). (B) Production of pulmonary surfactant in alveolar epithelial cells increased to a greater extent during air-liquid interface (ALI) culture than in submerged liquid culture. Surfactant production was quantitated by measuring the fluorescence intensity of lamellar body staining inside the alveolar epithelial cells (data represent the mean ± SEM of three separate experiments; *p < 0.001). (C) Physiological levels of cyclic strain did not increase the rate of FITC-albumin transport across the alveolar-capillary barrier. (D) Priming the alveolar-capillary barrier with cyclic stretch with 10% at 0.2 Hz strain did not cause any changes in the barrier permeability, regardless of its duration.

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Fig. S2. Cyclic strain-induced alignment of vascular endothelial cells. (A) Human umbilical vein endothelial (HUVE) cells cultured in the stretching microfluidic device for 10 hours without cyclic strain. (B) HUVE cells after 10-hr cyclic stretch with 10% strain at a frequency of 1 Hz. A non-porous PDMS membrane was used to clearly visualize morphological changes in the cells (see also Movie S4). (C) The orientation angle (θ) is defined as the angle between the major axis of the best-fit ellipse around a cell and the axis of cyclic strain. (D) Quantitative analysis shows a uniform distribution of the orientation angle in the absence of strain. (E) When the cells are subjected to cyclic stretch, the distribution of the orientation angle is skewed towards 90°, which represents the direction perpendicular to the applied strain. (F) Over 95% of the cells undergo reorientation and alignment in response to cyclic stretch, as compared to control without strain (*p < 0.001). Cells with orientation angles between 60° and 120° were considered aligned. Scale bars, 200 μm.

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Fig. S3. (A) Endothelial expression of ICAM-1 in response to stimulation of alveolar epithelial cells with TNF-α at 50 ng/ml. The cells were subjected to cyclic stretch during TNF exposure. Application of physiological cyclic strain (10% at 0.2 Hz) prior to or during epithelial exposure to TNF did not change the level of ICAM-1 expression in endothelial cells. Data represent the mean ± SEM obtained from two separate experiments. (B) After initial contact and rolling, a neutrophil (shown with an arrow) becomes activated and firmly adheres to an activated endothelium (not visible because it is unstained in this view) by spreading and increasing its contact with the endothelial surface within seconds after initial contact. Scale bar, 25 μm.

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Fig. S4. Epithelial stimulation with colloidal silica nanoparticles results in firm adhesion and infiltration of neutrophils from the microvascular canal into the alveolar microchannel. Yellow arrows show the neutrophils that accumulate on the apical side of the epithelium after transmigration through the alveolar-capillary barrier (not visible in this view). Scale bar, 40 μm.

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Fig. S5. (A) The strain-induced ROS response to silica nanoparticles is suppressed by 4 mM of NAC (time = 120 min). Data represent the mean ± SEM of three separate experiments; *p < 0.001. (B) Silica nanoparticles introduced into the alveolar air space cause a significant but delayed increase in the level of intracellular ROS in the underlying endothelial cells in the presence of physiological mechanical strain. As compared to the ROS response of the epithelium, endothelial cells increase their ROS production more gradually, and the maximum level of ROS achieved over 2 hours is lower. Data represent the mean ± SEM of three separate experiments.

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Fig. S6. ROS responses of alveolar epithelial cells to 16 nm quantum dots coated with carboxylic acids. The cells increase the level of intracellular ROS during the treatment with 50 nM of 16 nm carboxylated quantum dots while being stretched with 10% strain at a frequency of 0.2 Hz (square). Under these conditions, oxidative stress increases continuously over 2 hours. When exposed to quantum dots (triangle) or mechanical stretch (diamond) separately, the cells do not exhibit any significant ROS responses. ROS generation was normalized to the mean ROS value at time = 0 min; data represent the mean ± SEM of three separate experiments.

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Fig. S7. Intracellular ROS production in alveolar epithelial cells over 25 hours. Colloidal silica nanoparticles alone induce a gradual increase in oxidative stress over the period of 25 hours (triangle). After 20 hours, ROS generation reaches similar levels observed after 2-hr exposure to silica nanoparticles in the presence of physiological mechanical strain (square). Cyclic stretch of 10% strain alone (diamond), however, does not elicit ROS responses over 25 hours. ROS generation was normalized to the mean ROS value at time = 0 hr. Data represent the mean ± SEM of three separate experiments.

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Fig. S8. Alveolar epithelial cells cultured in our microfluidic device do not increase ROS generation when exposed to (A) single-walled carbon nanotubes (SWNTs) or (B) gold nanoparticles, even in the presence of mechanical strain. The SWNTs are 1 nm in diameter and 300 nm in length. The average diameter of the gold nanoparticles is 3 nm. ROS generation was normalized to the mean ROS value at time = 0 min. Data represent the mean ± SEM of three separate experiments.

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Fig. S9. (A) Representative confocal images and cross sectional views (a-d) illustrate increased uptake of fluorescent nanoparticles (magenta) into endothelial cells when they experience mechanical strain compared to cells cultured without strain. Internalized nanoparticles are indicated with arrows; blue and green represent nuclear and cytoplasmic staining, respectively. (B) The fraction of cells that internalize nanoparticles increased from ~ 7% to over 75%, due to application of 10% strain.

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Fig. S10. Intratracheal nebulization in an ex vivo mouse lung ventilation perfusion model delivers fluorescent nanoparticles to lung parenchyma. (A) Photomicrograph showing the tapered tip of Aeroprobe© Intracorporeal Nebulization catheter used for intratracheal nebulization in the mouse. (B-C) Photographs of the component parts and a steel tracheal tube (at the end on the right) (B) that are used to assemble the specialized catheter system which is fully assembled in (C). (D) Photograph showing positioning of the specialized nebulization catheter system and tracheal tube that are secured inside the mouse trachea during mechanical ventilation. Heart and lung are indicated with arrows. (E) Photomicrograph of tracheal tube, pulmonary artery (PA), and left atrial (LA) catheters during ex vivo lung ventilation perfusion. (F) Fluorescent micrograph showing density of 20 nm fluorescent nanoparticles used in the experiments on a glass slide after single pulse (5 ms, 100 psi) directed onto a glass surface. (G-H) Fluorescent micrograph of 5 μm lung section showing the density of same fluorescent particles seen in (F) on the luminal surface of mouse small bronchioles viewed in cross section (G) and sagittal sections (H) following nebulization and ex vivo ventilation perfusion for 60 minutes. Scale bars are 1 cm in (A-C), and 50 μm in (F-H).

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Fig. S11. Pre-conditioning of the endothelial layer in our microfluidic device with a shear stress of 15 dyne/cm2 resulted in an approximately 1.3-fold increase in the rate of nanoparticle translocation across the alveolar-capillary barrier. In contrast, lower levels of shear stress fail to produce similar results. The duration of pre-conditioning was 2 days. Translocation was normalized with respect to the rate of nanoparticle transport measured in the barrier primed with 0.1 dyne/cm2. Data represent the mean ± SEM of three separate experiments; *p < 0.05.

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Nanomaterials

Surface coating Size

ROS response (0% strain)

ROS response (10% strain)

Carboxyl groups 500 nm No No

Carboxyl groups 200 nm No No Polystyrene nanoparticles

Amine groups 200 nm No No

Carboxyl groups 16 nm No Yes Quantum dots

polyethylene glycol 13 nm No No

Silica nanoparticles N/A 12 nm No Yes

Magnetic iron nanoparticles

Carboxyl groups 50 nm No Yes

Gold nanoparticles N/A 3 nm No No

Table S1. Summary of cellular response to nanoparticulates in the lung mimic device. ROS response shown here indicates significant intracellular production of ROS observed over the course of 2 hours after the introduction of nanomaterials into the alveolar channel of the microfluidic system. Cells were treated with varying concentrations of nanomaterials; 100, 50, and 10 μg/ml for polystyrene nanoparticles, quantum dots, and silica nanoparticles; 100 and 20 μg/ml for magnetic nanoparticles; 10 and 1 nM for gold nanoparticles.

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References S1. S. Takayama et al., Adv. Mater. 13, 570 (2001). S2. M. I. Hermanns, R. E. Unger, K. Kehe, K. Peters, C. J. Kirkpatrick, Lab. Invest. 84,

736 (2004). S3. A. A. Birukova, A. Rios, K. G. Birukova, Exp. Cell Res. 314, 3466 (2008). S4. C. C. Dos Santos, A. S. Slutsky, J. Appl. Physiol. 89 (2000). S5. D. J. Tschumperlin, J. Oswari, S. S. Margulies, Am. J. Respir. Crit. Care Med. 162,

357 (2000). S6. M. Y. Liu, A. K. Tanswell, M. Post, Am. J. Physiol. Lung Cell Mol. Physiol. 277, L667

(1999). S7. D. J. Tschumperlin, S. S. Margulies Am. J. Physiol.-Lung Cell. Mol. Physiol. 275,

L1173 (1998). S8. B. Halliwell, M. Whiteman, Br. J. Pharmacol. 142, 231 (2004). S9. The ICAM1 antibody developed by Elizabeth Wayner/Gregory Vercellotti was

obtained from the Developmental studies Hybridoma Bank developed under the auspices of the NICHD and maintained by the University of Iowa, Department of Biological Sciences.

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Movie S1 shows real-time cyclic stretching of the alveolar-capillary bilayer formed on a porous PDMS membrane embedded in our microfluidic device. Cyclic mechanical strain was applied in the vertical direction in this view. Movie S2 shows real-time cyclic stretching of an array of fluorescent particles patterned on a PDMS membrane. Each dot is 10 μm in diameter and contains multiple 500 nm fluorescent polystyrene nanoparticles. Cyclic strain was applied in the vertical direction in this view, and the membrane cannot be seen in this movie. Movie S3 shows real-time cyclic stretching of human umbilical vein endothelial (HUVE) cells grown on a membrane in our device. A non-porous PDMS membrane was used to more clearly visualize mechanical stretching of the cells. Cyclic strain was applied in the vertical direction in this view. Movie S4 shows reorientation and alignment of HUVE cells in response to a cyclic stretch of 10% strain at 1 Hz over the period of 10 hours. A non-porous PDMS membrane was used to more clearly demonstrate morphological changes in the cells. Movie S5 shows fluorescently labeled human neutrophils flowing over a non-activated endothelium in the vascular microchannel. The alveolar-capillary barrier (not visible in this movie) was mechanically stretched with a cyclic strain of 10%. The neutrophils are observed to flow by the endothelium without adhering to the endothelial cells, as observed in normal vessels in vivo. The movie is played at 5 times real time. Movie S6 shows fluorescently labeled human neutrophils flowing over an endothelium activated by addition of TNF-α (50 ng/ml) to alveolar epithelial cells grown on the opposite side of the culture membrane. The alveolar-capillary barrier (not visible in this view) was stretched with a cyclic strain of 10%. The flowing neutrophils are captured by the activated endothelial layer and they attach to the cells even in the presence of continuous fluid flow and cyclic stretch, just as they do at sites of inflammation in vivo. The movie is played at 5 times real time. Movie S7 shows the entire process of neutrophil recruitment from the vascular channel to the alveolar chamber. A fluorescently-labeled neutrophil first adheres to the surface of the activated endothelium, crawls around, and then migrates into the subendothelial space and through a pentagonal pore on the membrane to cross the alveolar-capillary barrier where it goes out of focus. Note that the tissue layers are not visible in this view. The alveolar-capillary barrier was continuously stretched with a cyclic strain of 10%, but mechanical loading was switched off briefly at the time each frame was acquired to visualize neutrophil movement clearly without rocking motion generated by cyclic stretch. The movie is played at 5 times real time. Movie S8 shows phagocytosis of two attached E. coli (green) bacteria by a fluorescently-labeled human neutrophil (red) on the surface of the alveolar epithelium of the microfabricated lung mimetic device. The neutrophil, which was recruited to the alveolar epithelial surface after being stimulated to transmigrate across the barrier (not visible) by the presence of the bacterium, moves towards them and engulfs them. Cyclic stretch

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was turned off when each frame was captured; the movie is played at 300 times real time. Movie S9 shows another example of bacterial phagocytosis by fluorescently labeled human neutrophils (red) in the microfabricated lung mimetic system. The neutrophil that appears on the left (shown with an arrow) engulfs multiple bacteria within 5 minutes. The movie is played at 300 times real time.