At the Cusp of Biology and Materials - American Chemical Society

Review pubs.acs.org/journal/abseba

Advances and Challenges in Recapitulating Human Pulmonary Systems: At the Cusp of Biology and Materials Piyush Bajaj,†,‡ Jennifer F. Harris,§ Jen-Huang Huang,§ Pulak Nath,⊥ and Rashi Iyer*,† †

Information Systems and Modeling, §Bioscience Division, and ⊥Physics Division, Los Alamos National Laboratory, Los Alamos, New Mexico 87545, United States,

ABSTRACT: The aim of this review is to provide an overview of physiologically relevant microengineered lung-on-a-chip (LoC) platforms for a variety of different biomedical applications with emphasis on drug screening. First, a brief outline of lung anatomy and physiology is presented followed by discussion of the lung parenchyma and its extracellular matrix. Next, we point out the technical challenges in recapitulating the complexity of lung in conventional static two-dimensional microenvironments and the need for alternate lung platforms. The importance of scaling laws is also emphasized in designing these in vitro microengineered lung platforms. The review then discusses current LoC platforms that have been used for testing the efficacy of drugs or as model systems for investigating disorders of the lung parenchyma. Finally, the design parameters in developing an ideal physiologically relevant LoC platform are presented. As this emerging field of organ-on-a-chip can serve an alternative platform for animal testing of drugs or modeling human diseases in vitro, it has significant potential to impact the future of pharmaceutical research. KEYWORDS: drug screening, extracellular matrix, lung-on-a-chip, microengineered, toxicology

1. INTRODUCTION 1.1. Anatomy and Physiology of Human Lung. The very existence of human life is dependent on breathing and optimal gas exchange. This important function is served by a complex structure that provides for a large surface area for efficient gas exchange, the lungs. Lungs are large, spongy, airfilled organs located in the chest cavity protected by the ribcage. The right lung being the larger of the two has three lobes, whereas the left lung has two lobes, leaving space in the chest cavity for the heart.1 Although the primary function of the lungs is to provide the vital gas exchange of transporting oxygen into the blood and removal of CO2 from the blood required for our survival, the lungs also maintain blood pH, filter xenobiotics from the blood, and act as a blood reservoir. There are two main airway zones of the lung, the conducting zones and the respiratory zones. The conducting zones form a passageway for air to enter and exit the lungs, whereas the respiratory zones major function is gas exchange. Air enters the body through the conducting zones, passes through the pharynx and larynx into the lower airways: trachea, primary bronchi, bronchioles, and the terminal bronchioles. Air then © XXXX American Chemical Society

reaches the respiratory zone which includes respiratory bronchioles, alveolar ducts, and alveolar sacs (Figure 1). This progressive bifurcation of the lungs is addressed by their generation number and there are a total of 23 generations of airways in a normal human lung.2 Table 1 provides the approximate dimensions and the parameters of air velocity (U) and Reynolds number (Re) for each of the 23 generations of the human airway at about 3/4 the maximal inflation.1−3 Equations 1 and 2 were used for calculating the U and Re respectively.3 Un =

Ren =

Q 4Q = n 2 An 2 π dn ρUndn 4ρQ = n μ 2 μπdn

(1)

(2)

Received: November 11, 2015 Accepted: February 11, 2016

A

DOI: 10.1021/acsbiomaterials.5b00480 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 1. Dichotomy of the human lung. Human lung consists of a fractal structure of bronchioles and grapes like alveolar sacs.

Table 1. Human Airways Model of an Average Adult at 3/4 the Maximum Lung Inflation and the Important Parameters of U and Re1,2 (Table adapted with permission from ref 2. Copyright Springer.)

Q is the flow rate within the lung (4000 cm3 s−1),4,5 ρ is the density of humid air at 37 °C (1.2 × 10−6 kg cm−3),4,5 and μ is the viscosity of humid air at 37 °C (0.01958 cP).4 As the generation number increases (for n > fourth generation), the total cross-sectional area increases as well. Initially, the area increases slowly but as one reaches the respiratory zone, there is an exponential increase in the area. Re decreases as one goes deeper in the lungs, indicating that in the respiratory zone viscous forces dominate over inertial forces. Turbulent flow (Re > 4000) is seen in the first five generations of the lung airways (0−4), whereas laminar flow (Re < 2300) occurs from generation six onward.4 Transitional flow is seen in generation five where the Re is between 2300 and 4000.

Inspiration is an active process while expiration under resting conditions is a passive process where the lung returns to its equilibrium position. In order to breathe, the external intercostal muscles must contract, which moves the ribcage up and out. The diaphragm, most important muscle for inspiration, moves down by about 1 cm (normal breathing) at the same time, creating a small negative pressure (intrathoracic or intrapleural pressure, PIP) within the alveoli with respect to the atmospheric pressure (Patm = 760 mmHg). PIP is approximately 753 mmHg at the apex of the lung, whereas 758 mmHg near the base during inspiration with an average of around 756 mmHg. The gradient in PIP is the result of gravity and posture. The mechanics of breathing follows Boyle’s law. B


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ACS Biomaterials Science & Engineering Thus, as the pressure gets lower, air flows into the airways and volume increases.1 Expiration on the other hand mostly occurs as a result of the natural elasticity of the lungs which produces a positive pressure of +3 mmHg for a total pressure differential of 7 mmHg from inspiration to expiration,6 although healthy humans are able to force some air volume out of their lungs, which can affect this pressure differential. Lung elasticity is the natural ability of the connective tissues to recoil during expiration. In order to exhale, the lungs must return to a resting state. Positive internal pressure results from both inhalation and the relaxation of the diaphragmatic muscle. This causes air to naturally flow out if the elasticity of the lung is intact. Lung compliance (ratio of change in the lung volume to the change in the pleural pressure) is the ability of the alveoli and lung tissue to expand on inspiration. Compliance is reduced by chronic fibrotic diseases and increases in proteolytic diseases such as emphysema. Connective tissue is of utmost importance to compliance and elasticity of the lung. In turn, gas exchange is also reliant on compliance and elasticity due to the requirement of specific partial pressures for gas exchange to occur. Essential structural requirements for efficient gas exchange are total surface area, thickness, compliance, and elasticity of the tissue barrier and total pulmonary blood volume. The diffusing capacity of a gas exchanger correlates directly with the surface area and inversely with the thickness of the blood−water/gas (tissue) barrier.7 Epithelial cells form a continuum along the entire surface of the airways with the epithelium in the trachea and bronchi forming a pseudostratified, ciliated, columnar epithelium populated with ciliated, neuroendocrine, goblet, club, and basal cells. Along these conducting zones of the upper respiratory tract, the epithelium invaginates into numerous submucosal glands lined with serous and goblet cells.8 The height of the healthy epithelium in the trachea is roughly 25− 40 μm as measured from the basement membrane;9 this becomes progressively thinner and flatter the deeper one gets into the lung. In the smaller bronchioles, the cellular population of the airway epithelium changes such that the goblet cells are replaced by club cells10 with foci of neuroendocrine cells8 and the epithelium becomes cuboidal with progressively fewer ciliated cells. The terminal bronchioles divide to form respiratory bronchioles that contain alveoli and terminate in alveolar sacs, comprising thin-walled membranes where gas exchange occurs. At this point in the branching architecture of the lung, the respiratory zone begins. The alveolar sacs are lined on one side by alveolar type 1 cells, and on the other by vascular endothelium. Figure 2 shows a simplified version of the airway illustrating the distribution of various types of epithelial cells based on their location in the lung. Although the lung contains many additional cell types including immune system cells, smooth muscle, progenitor cells,8,11,12 and a complex lung vascular system,13 this diagram is meant to illustrate cells lining the airway epithelium, which are the cells that see the majority of pathogens or toxins first, and are thus the most common cell types sought after for LoC populations. The alveoli are supported by a thin basement membrane and lined with Alveolar type 1 (AT1) and Alveolar type 2 (AT2) cells on one side and the microvascular endothelium on the other (Figure 2). AT1 cells are extremely thin and flat and are primarily involved in gas exchange.14,15 They comprise only around 8−11% of the total cellular population in the parenchyma of the lung, yet cover around 90% of its surface

Figure 2. Simplified model of the predominant cell-types of the human lung. The simplistic model shows the distribution of various types of lung cells based on their locations in the lung. This model does not represent the full composition of native lung. The alveolar− capillary membrane, which is composed of alveolar epithelium, basement membrane, and microvascular endothelium, can also be seen.

area.16 On the other hand, AT2 cells make up around 12−16% of the cells in the lung parenchyma but cover only 7% of its surface.16 The primary function of these cells is production of surfactant that reduces surface tension at the air−liquid interface.14 These cells also act as “caretakers” of the alveolar compartment, quickly replenishing vulnerable AT1 and AT2 cells.14 There are close to 300 million alveoli in an average adult human lung with a combined alveolar surface area is 50−100 m2 for efficient gas exchange.1 Each of them is ringed by a capillary network.1,17 Alveoli are not seen until the respiratory zones and almost 50% of the total number of alveoli are present in the last generation (Table 1).2 Turbulent flow in the first five generations of the lung airways leads to chaotic mixing and deposition of microscopic dust particles in the upper conducting zone.18 This prevents them from traveling further and getting trapped in the lower airways which are critical for gas exchange. Additionally, the lungs have other natural protective mechanisms to prevent particles from entering the deep lung. Ciliated and mucusproducing cells in the nose, trachea, and bronchi act as a filter when breathing in, which prevents large particles from entering C


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healthy tissue physiology. Its components provide structural support for the growth of cells and for establishing cell−cell contacts. It is also responsible for a variety of other cellular functions such as migration, growth, fate determination, selfassembly for development of organs, and apoptosis.29 In addition, ECM is involved in wound healing by acting as a reservoir for growth factors and cytokines which are activated by injury.30 Thus, ECM is an essential contributor to organ integrity and tissue homeostasis. The pulmonary ECM is primarily composed of collagen I and III, which provides the necessary tensile strength, collagen IV and laminin which make up the bulk of the alveolar and basement membranes, and elastin which provides elasticity to the ECM.30 Other ECM proteins such as fibronectin and glycosaminoglycans (GAGs) are responsible for maintaining the polarity of the lung cells and their survival.30 Even though there are a number of proteins in the lung ECM, collagen is by far the most abundant and the main component of the pulmonary ECM.31 In addition, matrix metalloproteinases (MMPs), a family of zinc enzymes, are responsible for maintaining the continuous turnover (>10% per day) of these ECM molecules.31 The architectural organization of these ECM proteins together determines the overall properties of the pulmonary parenchyma in terms of its elastic and resistive (dissipative) behavior.32 This in turn directly influences cellular signaling, surfactant production, tissue remodeling, contraction of smooth muscle cells, and thus lung physiology.33 Normal native human lung has stiffness (Young’s modulus) of 1.96 ± 0.13 kPa; during different diseased states, this value can change considerably.30 During idiopathic pulmonary fibrosis (IPF), the stiffness of the lung significantly increases to 16.52 ± 2.25 kPa.30 IPF is the result of marked increase of ECM deposition in the pulmonary interstitium.34 As a result of this increased stiffness, the lungs cannot properly move oxygen in the bloodstream. Fibroblasts, main producer of pulmonary ECM, seeded onto acellular IPF showed increased α-smooth muscle actin (SMA) production compared to normal acellular lung characterized by low α-SMA production.30 This indicates that the pulmonary ECM may influence the cellular phenotype and the elastic modulus of the ECM may influence physiological properties of the lung tissue, though there is still much research needed to elucidate the full roles of each of

respiratory zones. The lining of the upper conducting zones contain mostly ciliated cells with a few mucous-producing cells. Mucin forms a thin coat on the surface of the ciliated cells and is swept up toward the oropharynx by the cilia. Foreign particles, pathogens and dead cells are trapped in the mucus and drawn up the mucociliary ladder and eliminated by swallowing and excretion. Figure 3A, B shows image of ciliated

Figure 3. Immunofluorescent images of human bronchial/tracheal epithelial cells after 21 days in air−liquid interface. Column 1 shows the cilia stained by β-tubulin antibody, column 2 shows the tight junctions visualized by actin staining, and column 3 shows the merged image. Top row shows the low-magnification image, scale bar = 100 μm. Bottom row shows the high-magnification image, scale bar = 25 μm.

normal human bronchial/tracheal epithelial cells after being differentiated for 21 days at the air−liquid interface. Lungs are also monitored closely by different types of immune cells such as neutrophils, dendritic cells, macrophages and lymphocytes which are responsible for its protective mechanisms. All of these immune cells work in cohort to modulate each other’s effect in the native lung to maintain homeostasis. 1.2. Extracellular Matrix of the Lung and Lung Pathophysiology. Physical properties of the cellular microenvironment and the extracellular matrix (ECM) play a key role in controlling the differentiation/functionality of cells, its pathophysiology, morphology, organ development and maintaining its functionality.19−28 ECM is a highly complex and dynamic microenvironment that is responsible for maintaining Table 2. Commonly Used Cell Lines in LoC Systems Today cell type bronchial epithelial (NHBE, HBTEC, HBEpC) alveolar type 1 alveolar type 2 small airway epithelial cells lung fibroblasts microvascular endothelial

A549 H441 immortalized bronchial epithelial (BEAS-2B, Nu-Li)

origin

application

commercial sources

available human primary cells usually derived from trachea and the primary bronchus on a chip,152 inflammation,153 conducting zone toxicology, goblet cell hyperplasia153 through secondary bronchi parenchymal-derived, sorted by alveolus on a chip,116 respiratory zone toxicology cell-surface markers parenchymal-derived, sorted by alveolus on a chip,116 respiratory zone toxicology, cell-surface markers surfactant studies isolated from the distal portion of the lung in mechanical injury,154 airway regeneration155 the ∼1 mm diameter airways whole lung-derived, sorted by recapitulation of tissue interface, ECM production152 cell-surface markers parenchymal-derived, sorted by recapitulation of tissue interface156 cell-surface markers frequently used immortal cell lines adenocarcinoma-derived mechanical stretch,156 ventilator injury157 adenocarcinoma-derived mechanical stretch156 derived from primary bronchus, bronchus on a chip, conducting zone SV40 or hTERT immortalized inflammation,158 goblet cell hyperplasia D

Lonza, Lifeline, Cell Applications, ATCC Sciencell Sciencell Lonza, Lifeline Lonza, Lifeline, Cell Applications Cell Applications

ATCC ATCC ATCC


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challenges. Human lung cells are generally harvested from excess tissues of surgical resections or lungs that are unsuitable or unmatched for transplant.58 Lungs unsuitable for transplant may include donors who perished from chest trauma or acute respiratory distress syndrome, which causes rapid breakdown of the cells. Healthy lung cells are challenging to obtain, and the healthiest cells generally come from nonsmoker lungs unmatched for transplant. Additionally, tissue donation and lung cell harvest must adhere to strict federal and international regulations (Title 21 Code of Federal Regulations, part 1271, subpart C)59 making it increasingly important to investigate other cell sources (for e.g., stem cells) which may be potentially used as a reliable source for lung cells. The use of primary bronchial epithelial cells (NHBE, normal human bronchial/tracheal epithelial) for the simulation of the bronchial tissue is well-established. These cells can be differentiated with highly specific media at the air−liquid interface to ciliated and mucin producing cells, among other cell types. There are several commercial sources (Lonza, Lifeline Cell Technology, Cambrex) and these cells have been successfully used to generate functional bronchial tissues in a planar format using Transwell inserts.60,61 Similarly, primary small airway epithelial cells (SAECs) have been differentiated to mimic the distal, small diameter airway cells. These cells have been used in toxicology studies of nanoparticles that reach the small airways and host−pathogen studies.62−65 Recently, mesenchymal stem cells (MSCs) were used to study lung injury and repair66,67 and recellularization of decellularized lung68,69 but techniques to grow functional tissue is still not mature. Other cell sources including epithelial progenitor cells, induced pluripotent stem cells (iPSCs), and embryonic stem cells70 currently have limited potential due to the difficulty in the reproducible differentiation of these cells into tissues using conventional 2D cultures and the limited knowledge of the optimal cellular microenvironment. New culture methods and approaches need to be developed to control the differentiation of these pluripotent stem cells. Platforms similar to the one developed by Hattori et al. can offer a high-throughput screening of the ideal lung microenvironment where they screened 16 unique cellular microenvironments for the culture of Chinese Hamster Ovary cells on a “micro-environment array chip”.71 These and other complex platforms could offer lung-specific microenvironments, enable tissue−tissue and tissue−ECM interactions and also dynamic lung-based biomechanical stimuli, which will induce these pluripotent stem cells to differentiate into functional lung tissue. Moreover, different types of lung cells have disparate differentiation media formulations, and the cells tend to dedifferentiate when exposed to alternative media. For example, bronchial epithelial tissues dedifferentiate to squamous cells when exposed to serum58 and vascular tissue generally requires serum, though recently, newer serum-free formulations have been developed for vascular systems.72 Alveolar tissue has been cultured in both serum-containing and serum-free media,73,74 though the main challenge to culturing AT1 and AT2 cells remains maintaining them in a differentiated state. Ideally, all the cell types of the lung need to be cultured in one single universal medium. This would provide optimal paracrine signaling opportunities between bronchioles, alveoli, vasculature, and the immune cells. However, there is currently no common media that can be used for all these cells. Thus, compartmentalizing cells in different chambers such that they

the molecules comprising lung ECM and the interaction between ECM and lung cellular behavior. For an in-depth discussion of the ECM and how it may play a role in repopulating decellularized lungs, see the review by Wagner et al.11 In asthmatic patients, remodeling of pulmonary ECM takes place which causes the muscles surrounding the lung to tighten up.35 This makes them narrower and increases the level of mucus production. Furthermore, all the components of the ECM such as collagen, elastin, and proteoglycans get altered in asthma and COPD.36 Almost 2−3-fold increased thickness of the basement membrane (subepithelial collagen) is seen in asthma.37 In emphysema, a type of COPD, there is imbalance of neutrophil elastase and MMPs, leading to the enzymatic degradation of elastin.38 As elastin is responsible for the maintenance of ECM elasticity, its degradation leads to increased stiffness of the ECM. Also, emphysema leads to the destruction of the alveolar air-sacs and as a result breathing requires more effort. 1.3. Challenges with Recapitulating the Complexity of the Lung. The use of relevant lung cells are one of the biggest challenges in developing a functional lung tissue in vitro. The human lung contains over 40 different cell types11 and multiple tissue types, comprising epithelium, stroma, endothelium, vascular tissue, immune tissue, smooth muscle, and cartilage. Recapitulating each one of these tissue types on a chip would require several cell types, not all of which are available commercially. In addition, the lung contains many progenitor cells that appear to have the ability to differentiate into multiple cell types. Many commercially available lung cell lines are derived from epithelial carcinomas. This may be due to the fact that most common cancerous tissue is epithelial in nature, and the first cell lines were tumor derived.39 Thus, the most common recapitulation of the lung is its epithelium. Table 2 provides the most commonly used primary and immortal cell lines in LoC systems today with descriptions and source. For an in-depth discussion of the lung cancer cell lines available, see the review by Gazdar et al.40 The most common source for in vitro lung studies are immortalized cell lines. For example, A549 (adenocarcinomaderived human alveolar basal epithelial) cells are widely used to study air−liquid interface,41 mechanical stretching,42,43 wound healing,44,45 reconstruct alveolar-capillary membranes,46 and regeneration of decelluarized lungs.47,48 Although A549 cells can also produce surfactant protein A, B, and C (reduce surface tension) and enhance gas exchange like primary human AT2 cells,49 literature reports conclude that the Raman spectra of A549 cells is statistically more similar to AT1 cells than AT2 cells.50 Early A549 studies concluded that the use of A549 cells as a model system for AT2 cells is questionable,51 whereas still other studies conclude that A549 cells are useful for modeling characteristics of AT2 pneumocytes.52−56 Nevertheless, studies in our laboratory (unpublished data) and others suggest that the functionality of A549 cells is not comparable to primary cells (i.e., poor cell−cell junctions).57 Therefore, although A549 cells might serve the purpose of some in vitro studies their use in the development of in vitro organ has not been optimized. The same holds true for immortalized bronchioepithelial cells such as BEAS2B, which are commonly used to mimic bronchial functionality. Although immortalized cell lines provide for a dependable source of lung cells, the use of primary human cells is highly desirable due to their ability to simulate in vivo responses with more fidelity. Nevertheless, primary cells present with their own E


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in investigating pulmonary disorders and evaluating the effect of drugs on human tissue samples. 1.5. Advantages of Lung-on-a-Chip Platforms and Their Biomedical Applications. Modern drug-discovery is a tedious and expensive process requiring a series of complex steps.85,86 The average length of time from target discovery to approval of a drug currently averages 12−15 years.86 Of all the drugs that make it to the Phase I trials, around 80% fail during development.86 The potential cause of this high rate of failure is the lack of human relevant testing models. Currently, animal models are used for drug testing and therapeutic validation. However, animal models often do not recapitulate human responses, as a result, compounds that are cleared in Phase I trials, sometimes fail in Phase II or III trials.85 Accounting for all the failures, the cost of a successful drug today can be in excess of $1 billion.87,88 Also, the number of new drugs approved by the U.S. Food and Drug Administration (FDA) per billion dollars spend in R&D by the pharmaceutical companies has approximately halved every 9 years since 1950 (inflation adjusted), a trend commonly called “Eroom’s Law” (an inverse contrast to the familiar “Moore’s Law”).89 Thus, fewer drugs are approved and the cost for approving them is getting higher. Therefore, there exists a clear need to test the drugs on platforms, which more closely represent human physiology and show a closer approximation of the native function for faster, cheaper, and more reliable drug discovery.90,91 OoC technology might hold the promise to shorten the screening process for drugs by providing an alternate pipeline and offering a physiologically relevant model system for humans. These OoC technologies can address the formidable pharmacological and physiological gaps between monolayer cell cultures, animal models, and humans.92 It might be possible that these OoC technologies can supplement or even completely replace part of the early clinical trials by enabling a variety of diseased and healthy tissue models for rapidly testing the efficacy of new drugs, tremendously reducing the cost and time required for drug development.82,93 Modulation of the cellular microenvironment in the lung-ona-chip (LoC) platforms has the potential to mimic diseasestates for investigation of pulmonary disorders and effective therapeutics. These LoC models will allow studying the dynamic response of drug/irritant to humans. For example: by introducing cigarette smoke in these LoC models, we might be able to study its effect on cilia beat frequency and mucus production in a dynamic environment similar to one experienced by lung cells in vivo. Static 2D cultures and animal models do not offer such a close replication of the native human functions for these studies. Cystic fibrosis (CF) and other types of genetic disorders can also be studied using these LoC models. By using iPSCs from patients suffering from CF and using the complex architecture of a LoC model, it might be possible to closely approximate the in vivo conditions experienced by the cells in their native environment and develop better drugs to counter it. Mucin plugs and mucous cell metaplasia are common in disease states such as chronic obstructive pulmonary disease (COPD) and asthma,94,95 so failure to wash the mucin out of the airways may be an intentional way to recreate these disease states in a LoC model. The natural protective system could be recreated by periodically washing the epithelium. By changing the elasticity or compliance of the substrate, it might be possible to study diseases like IPF,30 emphysema,33 and other disorders of the lung parenchyma.75 Culturing mutant AT2 cells that lack the

are connected to each other via a common surrogate medium with additions of specific factors in these chambers can be a solution until a common universal medium capable of supporting all cell types of the lung is available. As the bronchioles and the alveoli have very different geometrical features, culturing them in two-dimensional (2D) flat tissue culture dishes does not help to capture the characteristics of in vivo lung. It is well-known in biology that function is derived from the structure of the tissue. Thus, it is important to culture these cells in environments which closely mimic the geometrical features of their native counterpart. The rigid nature of conventional 2D platforms cannot mimic the soft in vivo lung microenvironment. Furthermore, the lung tissue is always under a pre-existing tensile stress (prestress) because of the ballooning effect caused by the transpulmonary pressure.75 The act of breathing causes cyclic stresses at the pleural surface of the lung parenchyma which gets transmitted via the ECM to the cells. Also, these static in vitro platforms are generally supplied with excessive amounts of nutrients and cannot generate the dynamic biochemical and mechanical stimuli, which are needed for the normal functioning of cells. Furthermore, standard in vitro reductionist approaches by coating one type of ECM on rigid tissue culture substrates do not truly account for the complex 3D lung-specific ECM structure and composition that are actually responsible for the intricate signaling cascades of the pulmonary system. Only by developing platforms that combine the ECM composition, geometry, topography, dimensionality and the dynamic nature of the lung tissue would it be possible to develop the right conditions to simulate and model the lung tissue in vitro. Having said that, it is no trivial task to simulate with any accuracy the simplest of lung tissues, let alone the sophisticated architecture that allows for its unique functions. Evolutionary pressure has guided the development of the lung as we know it, with the principle goal of optimizing gas exchange and the acquisition of oxygen. Indeed, what has taken nature millions of years to perfect can hardly be expected to be duplicated by man within a span of a lifetime. Therefore, although one might aspire to simulate this highly complex and sophisticated paradigm, we must be satisfied, at least for now, to discuss possibilities of mimicry, albeit with much less accuracy. 1.4. Alternative Platforms for Recapitulating the Complexity of Lungs. Advances in microfluidics and microfabrication have now enabled researchers to design relatively complex tissues on a chip.76,77 Some of the challenges discussed in the previous section can be addressed by using these novel in vitro tissuelike microsystems termed as “organon-a-chip (OoC)”. By combining biocompatible materials, microfluidics, microfabrication and engineered control systems, it is possible to capture both the anatomy and physiology of the respiratory system in a time-varying dynamic platform. Simultaneously, these OoC systems can also address the physics and physiological constraints of a human lung and the diversity of microenvironments present in it. These microstructures recapitulate physical microenvironments such that when mammalian cells are subjected to the physical forces imposed by the structure coupled with fluidics and growth factors from the media encourages differentiation into tissues which perform organlike functions. Current studies suggest that modern OoC respond by producing proteins and tissue like structures that would otherwise not be possible in a conventional static 2D culture of primary cells.78−84 Thus, these OoC platforms have the potential to accelerate research F


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cells within them. For a millilung, it will not be possible to fabricate a device based on the dimensions seen in Table 4 while maintaining a physiologically relevant lung blood (culture medium) volume. By reducing the lung blood volume to less than 10% of the systemic total blood (culture medium) volume, a device of width 70 000 mm (7 m) would be required, which is far larger than the size of a silicon wafer for a milli-lung while for a microlung, the compartment volume would be less than 1 μL, preventing cell growth.98 In addition, the height of the device (milli or micro) would be too small (4−6 μm) to allow the growth of cells, height of a pseudostratified epithelium is around 25−40 μm. Furthermore, very powerful pumps would be required to transport cell culture medium through devices of such dimensions. Similar problems are associated with the microlung as well. Also, it is not ideal to classify organs as purely 2D or 3D, as many aspects of an F-2D organs might be best scaled under the F-3D assumption.98 For example, modeling the growth of lung cancer on a LoC should be F3D as the growth of the fibroblasts would be constrained by volume and not surface area.98 As it would be challenging to use these allometric scaling laws for modeling the in vitro lung platform, one can use metabolic scaling and engineer alternative strategies such as culturing cells in resource-limited settings, which are transport accessible to mimic the in vivo tissue conditions.98 Reviews by Moraes et al.98 and Wikswo et al.100 provide a much more in-depth analysis of how scaling can be applied to LoCs or OoCs in general.

ability to produce surfactants will lead to high surface tension at the air−liquid interface. This can be used as a model system to study infant respiratory distress syndrome. These LoC platforms along with advances in stem cell technology will enable the realization of personalized medicine, drugs tailored toward individual patients. Thus, these LoC (OoC) devices have the potential to address a multitude of unmet needs of modern day drug discovery.

2. IMPORTANCE OF SCALING OoC technologies have advanced considerably in the past decade. However, understanding of biological scaling laws and how they apply to OoCs has been largely ignored. To replicate human physiology and drug response with human OoCs and the larger human organ constructs (HoCs), it is critical that the OoC/HoC has the correct relative size. Shuler and colleagues were the first to design chamber sizes and flow rates based on the size/mass of the organ and residence times.92,96−98 They concluded that a linear relationship exists between the organ and organism mass. Organs are classified as functionally twodimensional (F-2D) or functionally three-dimensional (F-3D); lungs, being constrained by the surface area, were classified as F-2D.98 The allometric scaling of different system components is described by eq 3 which shows the dependence of the biological variable Y on M with b as the scaling constant and Y0 is a constant that is characteristic of the organism.99 Y = Y0M b

3. CURRENT LOC PLATFORMS Microscale technologies initially developed for the electronics and the semiconductor industry have allowed researchers to create the complex cellular microenvironment and present to the cells biological, mechanical, and chemical cues in a physiologically relevant niche.76,77 Using the tools of softlithography, microfluidics, and microfabrication, researchers have mimicked a variety of complex organ systems like brain, 101,102 kidney, 103,104 gut, 81,105 heart, 84,106 and liver.107−109 In this section, we shall discuss the state of the art in the development of LoC which requires incorporation of multiple cell-types in the same platform while simultaneously providing a dynamic environment. The most advanced LoC platform to date was fabricated by Huh, Ingber et al., and the device demonstrated for the firsttime organ-level responses to bacteria and inflammatory cytokines.83 The device mimicked the alveolar-capillary interface which is one of the many functional units of the lung. Human alveolar epithelial cells (NCI H441, A549, and E10− immortalized cell lines) and human pulmonary microvascular endothelial cells were seeded on apical and basal sides of a 10 μm thin polydimethylsiloxane (PDMS) membrane coated with collagen and fibronectin, respectively. After achieving cellular confluence, air was introduced in the epithelial compartment of the device to mimic the air−liquid interface of the alveoli. The normal breathing cycle of the lung, was mimicked by applying a vacuum in the lateral microchambers of the device (Figure 4A). This resulted in the stretching of the thin PDMS membrane and provided for similar mechanical cues possibly experienced by cells in vivo. Using this platform, they showed that the stimulation of the epithelial surface with tumor necrosis factoralpha (TNF-α) increased the expression of the leukocyte adhesion molecule, intercellular adhesion molecule-1(ICAM-1) on the endothelial surface (Figure 4B). The activated endothelium promoted the adhesion of fluorescently labeled

(3)

Using Table 3 and lung design constraints of physiological shear stress (τ = 15 dyn cm−2)83 and the surface area (in vivo Table 3. Allometric Scaling of Different System Components Using the Allometric Scaling Lawa

a Table adapted with permissiom from ref 98. Copyright 2013 The Royal Society of Chemistry.

surface area of lung = 70 m2),98 Table 4 is generated, which shows a parametric sweep of possible lung design configurations for a milli (1 × 10−3) and micro (1 × 10−6) LoC along with the important parameters of U and Re for each of those device configurations.3,98 The height (h), U, and Re of the devices were calculated using eqs 1, 2, and 4. τ=

6μQ wh2

(4)

where τ is the shear stress, μ is the viscosity of the medium (1.11 cP),28 Q is the flow rate (cardiac output), w is the width of the device, A is the cross-sectional area of the device, and d is the characteristic dimension (rectangular tube d = 2wh/(w +h)).3 It can be clearly seen that allometric scaling cannot be employed for a milli or microlung as it will create some physical challenges in the fabrication of the device or for culturing the G


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Table 4. Parametric Sweep of Possible Lung Configurations and the Corresponding U and Re for a Millilung and a Microlunga

a

Table adapted with permission from ref 98. Copyright 2013 The Royal Society of Chemistry.

junctions. Finally, a newly developed transient receptor potential cation channel subfamily V member 4 (TRPV4) channel blocker pharmacological agent, GSK2193874 (GlaxoSmithKline) completely inhibited the leakage of fluid induced by IL-2. As the beneficial effects of GSK2193874 are yet to be confirmed in humans but are validated in animal models,111 this platform clearly showed the potential of OoC platforms for toxicology, pharmacology, and drug efficacy. A previous generation of the above device was used by Huh and coworkers to mimic the impact of pathologic liquid flow plugs seen in the respiratory system on the small airway epithelial cells.112 They showed that the rupture of liquid plugs that simulates surfactant deficiency led to the injury of the epithelial cells because of the mechanical stresses generated by the rupture. This microfluidic OoC construct was useful in deconvoluting the impact of complex mechanical forces on the cells and could be used for understanding lung injuries which results from fluid stresses. Another excellent example on the gain made by these platforms over static monolayer cells culture systems is the amenability of these devices for the understanding of mechanical stress on lung performance. Takayama and colleagues fabricated a microfluidic alveolar epithelial cellculture platform where the pathophysiological effects of both the solid and fluid stresses could be examined individually and in combination.43 Mechanical ventilation is an important therapy for patients suffering from acute respiratory distress syndrome (ARDS). However, it often results in acute pulmonary cellular injury termed as ventilator induced lung injury (VILI).113 VILI is the combination of solid mechanical stresses (cyclic stretch of alveolar epithelial cells) and fluid mechanical stresses (shear gradients over the cells). However, until recently, current in vitro platforms have been unable to examine the effect of each of these stresses in combination on epithelial cells. The Takayama microfabricated device consisted of three different components shown in Figure 5A: (1) chamber for culturing alveolar A549 cells, (2) PDMS membrane for mechanical stretching, and (3) actuation channel for controlling the stretch and meniscus propagation. By using 35−40% cyclic strain and 3.44 mm s−1 fluid meniscus velocity, they were able to simulate the Re for the 17th generation of the bronchioles. Completely filling the culture chamber with media established an environment where the cells on the PDMS membrane experienced cyclic mechanical stress but minimal fluid stress. With no fluid in the culture chamber, the cells on the membrane only experience solid mechanical stress and no

Figure 4. Human breathing LoC microdevice. (A) Mechanical stretching of PDMS membrane driven by vacuum can recreate physiological breathing movements. Reconstitution of inflammation and infection responses in LoC device (b, c). (B) Epithelium was simulated by TNF-α in the device. (C) Human neutrophils were adhered to activated endothelium. Reproduced with permission from ref 83. Copyright 2010 The American Association for the Advancement of Science.

human neutrophils. This phenomenon was not observed in the nonactivated endothelium. More importantly, the adhered neutrophils demonstrated extravasation (diapedesis) in response to the bacteria present on the alveolar surface and engulfed them in a matter of minutes (Figure 4C). This microfabricated in vitro LoC system was thus able to mimic the immune response to microbial pathogens as seen in in vivo human lungs. They also demonstrated the expression of ICAM1 in response to the exposure of the alveolar epithelial cells to silica nanoparticles. Enhanced levels of ICAM-1 expression was recorded when the system simulated normal breathing cycles by the application of 10% cyclic strain at 0.2 Hz to the thin PDMS membrane. Finally, they also showed that the addition of the nanoparticles to the alveolar surface lead to the production of reactive oxygen species, which was otherwise not seen without stimulation. Conventional static cultures are unable to emulate the complexity of responses as is possible by using this in vitro biomimetic LoC platform. This platform represented an innovative low-cost system with potential application of supplementing or even replacing animal studies and in vivo assays for pharmacology/toxicology studies. Another advantage of using such systems is their ability to simulate diseased conditions. The same group modeled the diseased state of pulmonary edema and used pharmacological agents to treat it.110 Addition of interleukin-2 (IL-2) to the vascular endothelium channel resulted in the leakage of fluid into the alveolar epithelial compartment and this leakage worsened even further on application of cyclic stretches. Reversal of this pulmonary edema was possible by the addition of angiopoietin-1 which stabilized the vascular endothelial cell H


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support the growth of all the three primary cells, the researchers showed mucociliary differentiation and barrier function studies using their biomimetic lung microfluidic platform. Figure 6 shows the triple coculture of the differentiated primary airway cells in their microfluidic device.

Figure 5. Mechanical stress studies in microfluidic alveolar model. (A) Cross-sectional view of the microfluidic chip for inducing mechanical stress on A549 cells (B) Live/Dead image showing the impact of mechanical stress on the A549 cells after different number of stretch cycles. Red, dead cells; green, live cells. Reproduced with permission from ref 43. Copyright 2011 The Royal Society of Chemistry.

Figure 6. Triple coculture of human primary lung cells in a LoC device. (A) Schematic of the lung structure. (B) Schematic of the microfluidic chip. Immunofluorescence images of (C) airway epithelial cells, AE; (D) fibroblasts, Fb; (E) microvascular endothelial cells, MvE. Blue, Hoechst nuclear stain; green, phalloidin; red, mucin. Scale bar 50 μm. Reproduced with permission from ref 115. Copyright 2014 The Royal Society of Chemistry.

fluid mechanical stress. By partially filing the channel with fluid, the cells experience a combination of fluid and solid mechanical stresses because the fluid meniscus propagates over the cells during mechanical stretching. Thus, by varying the amount of liquid in the device, they were able to control the type of mechanical stress experienced by the cell on the 100 μm thin PDMS device. After 60 cycles of stretch, the cell monolayer experiencing both the solid and fluid mechanical stresses showed a large degree of detachment while it was intact in populations that were exposed only to solid stresses. Figure 5B is the live/dead fluorescent micrograph of the cells which experienced both the solid and fluid mechanical stresses. Their device enabled the investigation of mechanical stress on alveolar cells, hitherto not possible in current in in vitro systems. Xu et al., designed a microfluidic chip for the threedimensional (3D) coculture of lung cancer and stromal cells, mimicking the microenvironment found in lung cancer.114 The researchers used this drug-sensitivity platform for screening appropriate chemotherapy schemes for eight different patients in a dose-dependent manner and thus demonstrated the use of these in vitro microengineered lung platforms for personalized treatment in lung cancer. Although previous studies have used secondary or immortalized cell lines in LoC platforms to investigate normal or diseased pulmonary physiology, recently Sellgren et al., fabricated a biomimetic microfluidic device of the human airways using all primary cells.115 The researchers did a triple coculture of lung epithelial cells, fibroblasts and lung microvascular endothelial cells in different layers (compartments) to closely recapitulate the in vivo biology of the human lung. By using a gravity fed microfluidic system and culture medium to

Recently, Stucki et al., fabricated a unique LoC array capable of generating 3D cyclic strains mimicking the breathing movement. The device seeded with primary alveolar epithelial cells and subjected to these cyclic strains showed enhanced enzyme activity and improved transporter function, clearly demonstrating the importance of mechanical strain and the applicability of the device in pulmonary research.116

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PARAMETERS IN DEVELOPING AN IDEAL LOC PLATFORM The highly complex architecture of any human organ poses extreme difficulties in engineering an exact biomimetic model. The task is further complicated by design constraints imposed by the scaling law to develop OoC platform. The human lung operates under unique conditions dealing with two fluidic phases (air and blood) and is composed of two distinct regions (the bronchioles and the alveolar sacs). Further engineering challenges arise from the constant stretching (inflation and deflation) of the alveolar sacs. Different types of cells and ECM are present at different regions with very specific functions (e.g., surfactant production, elasticity, foreign particles/organisms migration, and gas exchange). Nevertheless, as discussed in the previous sections, many engineering solutions have been successfully investigated to closely capture many physiological and functional aspects of a healthy/diseased working lung. Although, the above-described LoC platforms have made significant progress in trying to capture the salient features of the native lung, most of these platforms have been proof-ofconcept studies and cannot yet be used for drug efficacy testing. By combining these platforms with iPSCs and optimizing the important engineering parameters, it would be possible to I


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membranes have been used to recreate the alveolocapillary barrier to understand pathogenesis and recovery after acute lung injury57 and to investigate the inflammatory and cytotoxic responses of the alveolocapillary model to silica nanoparticles.133 Although biocompatible, these membranes do not have the elastic properties of PDMS membrane and will not allow for changes in membrane geometry. Other properties of the membrane such as stiffness,21,22,27,134 geometry,19,135−137 topography,20,138−140 and pore size,141,142 also need to be closely monitored to mimic the elements of the in vivo lung. Hydrogels (synthetic or native or their combination) can be used as an alternative building material for these LoC platforms. By modulating the molecular weight, cross-linker concentration, and the degree of polymerization, these hydrogels may be better suited to mimic the soft microenvironments of the lung tissue and thus allow optimal differentiation of primary lung cells or stem cells to lung-specific cells. Although both synthetic and native hydrogels can be tailored to match the properties of the parenchymal tissue, native hydrogels in particular offers added advantages of having biologically active moieties where the cells can attach to and proliferate. Most synthetic hydrogels such as polyethylene (glycol) diacrylate, polyethylene (glycol) monoacrylate, polyacrylamide, etc., need to be chemically modified to allow cell attachment. On the other hand, most native hydrogels such as collagen, fibrin, gelatin, Matrigel, etc. readily allow cell attachment and growth. Additionally, native hydrogels can also be enzymatically degraded by the cells unlike synthetic hydrogels thus making them even more attractive. For an in-depth discussion on the advantages and disadvantages of both native and synthetic hydrogel, read the review by Bajaj et al.144 3D printing technologies such as stereolithography which allows mask-free fabrication of different polymers into mechanically tuned hydrogels would be an ideal way to fabricate different components of the LoC platforms and to precisely organize the cellular components of the lung. This and other 3D fabrication methodologies will allow the fabrication of LoC platforms in high-throughput fashion, thus bringing these proof-of-concept devices into the pharmaceutical industry. Certain LoC platforms have used uniaxial stretch to simulate the breathing motion of the lungs.83 While this unidirectional stretching demonstrates the effect of mechanical stress in these LoC platforms, it does not truly mimic the inflation/deflation of alveolar sacs. To closely model that, it would be important to capture the geometric and mechanical constraints of these fundamental units of gas-exchange. This can be achieved by drilling an array of circular holes in the substrate and using sinusoidal vacuuming to mimic the breathing cycle of the lungs, similar to Figure 5A. Application of vacuum will result in the distension of these membranes into half-balloon like structures much akin to alveolar sacs. This will allow for the lung epithelial cells to experience similar cyclic circular stress profiles as seen in the native lungs. For optimal representation of the alveolar system a coculture of AT1 and AT2 cells should be a prerequisite on the apical side of the scaffold with a ratio of the AT1 cells to AT2 cells similar to that seen for in vivo lungs to enable desirable concentration of surfactants.16 As AT2 cells spontaneously differentiate to AT1 cells,143 it would be important to prevent this by using biomaterials and advances in cell biology designed to maintain AT2 characteristics. Commercial sources of AT2 cells have proven to be unreliable in our hands and rapidly transform into non-AT2 cells, both morphologically and

develop a device that could be useful for future absorption, distribution, metabolism, and excretion (ADME) studies and pharmacokinetic/pharmacodynamics drug modeling. Figure 7

Figure 7. Schematic of a possible lung platform. Bioinspired LoC platform is comprised of both fractal bronchioles and bubble like alveolar sacs. Porous hollow fibers are developed to represent terminal bronchioles and part of respiratory bronchioles followed by scaling law. Each hollow fiber can grow ciliated and mucin producing bronchiolar cells to mimic the physiologically similar cellular microenvironment. One way in-and-out design allows lung platform to breath with controllable inhalation and exhalation cycles. Submicron thickness of elastic membrane is used to fabricate alveolar sacs directly connected to terminal/respiratory bronchioles. These alveolar sacs can tolerate the continuous inflation and deflation driven by pneumatic pumps. All bronchiolar and alveolar units are submerged in continuous flow of medium to maintain their physiological functions.

shows a possible device design. In this section, we discuss some of the design parameters that can be optimized in order to develop a closer approximation of the native lung in an in vitro LoC platform. A typical LoC platform consists of different compartments and are generally fabricated using PDMS.76,83,115 PDMS offers great flexibility in terms of fabrication of complex microstructures,117 rapid prototyping,118 facile bonding,119 elastic components,120 gas permeability,121 and optical transparency.122 However, its hydrophobic nature allows diffusion of hydrophobic small molecules within its walls and this can lead to difficulties in elucidating the correct metabolic responses from LoC platforms.123,124 This makes PDMS potentially challenging for toxicology studies. To avoid small molecule absorption, different surface modification techniques such as a sol gel treatment with silica nanoparticles,125 or grafting with hydrophilic polymers126,127 has been used. Extremely thin biocompatible membranes (∼0.4 μm)16 are required to mimic the function of the in vivo basement membranes, however, the very thin dimensions required to simulate the alveolar basement membrane make it handling difficult. Therefore, membranes that can be deposited in situ would greatly reduce the need for handling of the membranes. For example, parylene is a biocompatible polymer that can be vapor deposited and patterned using shadow masks or reactive ion etching.128−130 It has also been shown that mesh supported parylene membranes are suitable for adherence and proliferation of retinal pigment epithelial (RPE) cells.131 Parylene is also sufficiently elastic to facilitate cyclic inflation and deflation of the membranes.132 However, such novel materials are minimally investigated for application in LoC and may be considered for further investigations. Biocompatible membranes made of polyester have also been used for the culture and differentiation of normal human bronchial epithelial cells and to study drug transportation through the cell monolayer.60 Polycarbonate J


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systems engineering approach to control the different functions of the integrated bronchiolar and alveolar platform. Although a wide range of miniaturized ventilations systems, pumps, and valves are available, innovative solutions may be required to simplify the overall systems engineering. For example, recently, Whitesides and Takayama group demonstrated a feasible way to control the inflation of 64 PDMS balloons simultaneously over Braille pins using pneumatics.151 Similar systems can potentially be used for high-throughput inflation studies on a LoC platform. The monitoring system of the LoC does not necessarily have to be directly connected to the lung platform but should exist to validate the lung model system. Inspite of the technical challenges, which can potentially be resolved by some clever engineering and materials, the advantages of organ platform technologies far outweigh the drawbacks. These chip technologies present an opportunity to simplify drug discovery and fundamental research while simultaneously offering an unprecedented level of physiological complexity in in vitro systems. Multiorgan chips can further push the state-of-the-art in simulating human responses while allowing for pharmacokinetics and pharmacodynamic studies not possible by simple in vitro cultures. However, end users such as pharmaceutical companies, defense agencies (DARPA, DTRA, DOD) and regulatory agencies (NIOSH, OSHA) have to be convinced of the utility of these systems. All of these “customers” have a vested interest in the success of this technology due to the huge advantages they offer for drug screening and discovery, repurposing of failed drugs, rapid screening for medical countermeasures and environmental stewardship. In addition, acceptance of these systems as possible replacement or reduced use of animal models has to have “buy-in” from regulatory agencies such as the FDA. To gain the confidence of such agencies, these models will have to undergo significant validation using drugs with well-known toxicological profiles, ideally drugs that have passed animal testing but have failed in clinical studies. These platforms have to withstand stringent validation ideally by comparison to in vivo tissues/organs, but the ease of use of these systems is also critical. User-friendly interfaces and modularity of systems will be key in acceptance of these models by a broader but relevant audience.

functionally. This represents a significant challenge in the functional simulation of an alveolar compartment. In addition, a 3D microvascular tissue architecture is desirable to represent the in vivo environment of the native lung. However, growing microvascular cells in 3D on the baso-lateral side of the membrane is extremely challenging and may pose diffusion limitations on the apical side. Microvascular cells would need to be encapsulated in 3D porous hydrogels, which could be fabricated on the membrane itself using 3D printers or other similar 3D fabrication modalities.144 Generally, in the development of LoC platforms, the bronchiolar compartment is neglected mainly because of the complexity in its fabrication using current technologies. The bronchiolar unit consists of a 3D fractal structure which bifurcates into progressively smaller units (Figure 1). The scaffold used for the culture of bronchiolar epithelial cells should be hollow for the flow of air and also have pores to allow the exchange of gases and cell culture medium. Boucher’s group pioneered the culture of human airway bronchiolar epithelial cells on both the planar and cylindrical surfaces. They used these surfaces to study cystic fibrosis,145 transport characteristics of periciliary liquid (PCL) and mucus,146 and volume transport of airway−surface liquid.147 They found that the epithelial cells in the cylindrical biofiber system were columnar as seen for planar air−liquid interfaces but lacked (less than 5%) the number of ciliated cells. However, no clear hypothesis was presented for this low number of ciliated cells. Therefore, much research needs to be done in order to understand the growth and differentiation of bronchial epithelial cells in cylindrical hollow fiber systems. Understanding the mechanism of differentiation of epithelial cells in cylindrical geometries and establishing a long-term bronchiolar culture would greatly aid the research in primary bronchiolar disorders and parenchymal disorders with prominent bronchiolar involvement.148 Finally, to develop a complete model of a LoC system, integration of the alveolar and the bronchiolar unit would be required. Growing the two different types of tissue in the same culture medium will be challenging. Hickman’s group has developed serum-free cell culture medium formulation that enables the culture of neurons and muscles for extended periods of time.149 This same medium has also been utilized for other cell types like glial cells and cardiomyocytes as well.150 Similar medium formulations need to be developed to enable the growth and culture of alveolar and bronchiolar cells in the same microfabricated device. Currently, no cell culture medium exists that would allow the growth and differentiation of these two different cell types. In addition, the differentiation of the two cell types occurs at different time scales; therefore, strategies need to be developed to allow the differentiation of the two cell types. One approach would be to use a modular system, grow and differentiate both cell types individually, and then couple the two modules together prior to toxicity or pharmacologic analysis. Control system of the lung organ platform should consist of the following three parts: (1) a ventilation system which can generate periodic flow in both the forward and the reverse directions to support the inflation and deflation of the alveolar sacs; (2) a circulating system that pumps cell culture medium to allow the growth and differentiation of the cells; (3) a monitoring and evaluation system which can in real time determine the functionality of the parenchymal tissue by first testing it for gas exchange and then evaluating the response of the lung model to toxins. Such integration would require a

4. CONCLUSIONS This review summarizes some of the design challenges, scaling laws and strategies for developing LoC platforms. However, the field of OoC or more specifically LoC platforms is very much still in its infancy. Many biological, engineering, and fabrication related challenges need to be addressed before developing a LoC model that is capable of performing pharmacokinetic, pharmacodynamic and toxicology analysis. Nevertheless, given the rising costs of drugs and healthcare in general, there is an immediate need for such LoC platforms. Most of the current research on LoC technologies is being executed in academia, government laboratories and other nonprofit organizations. In order for this technology to gain a foothold, industrial buy-in and engagement by regulatory agencies is critical considering the far-reaching implications for the pharmaceutical industries and healthcare in general.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Telephone: 505-500-2486. Fax: 505665-5283. K


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(15) Desai, T. J.; Brownfield, D. G.; Krasnow, M. A. Alveolar progenitor and stem cells in lung development, renewal and cancer. Nature 2014, 507, 190 http://www.nature.com/nature/journal/vaop/ ncurrent/abs/nature12930.html#supplementary-information,. (16) Bonner, J. C.Respiratory Toxicology, In Molecular and Biochemical Toxicology, 4th edition, Smart, R. C.; Hodgson, E., Eds., 2008. John Wiley & Sons, Inc.: Hoboken, NJ, USA. DOI: 10.1002/ 9780470285251.ch27 (17) Weibel, E. R. What makes a good lung? Swiss Med. Wkly 2009, 139 (27−28), 375−86. (18) Radhakrishnan, H.; Kassinos, S. CFD modeling of turbulent flow and particle deposition in human lungs. Conf. Proc. IEEE Eng. Med. Biol. Soc. 2009, 2867−2870. (19) Bajaj, P.; Reddy, B.; Millet, L.; Wei, C.; Zorlutuna, P.; Bao, G.; Bashir, R. Patterning the differentiation of C2C12 skeletal myoblasts. Integrative Biology 2011, 3 (9), 897−909. (20) Bajaj, P.; Rivera, J. A.; Marchwiany, D.; Solovyeva, V.; Bashir, R. Graphene-Based Patterning and Differentiation of C2C12 Myoblasts. Adv. Healthcare Mater. 2014, 3, 995. (21) Bajaj, P.; Tang, X.; Saif, T. A.; Bashir, R. Stiffness of the substrate influences the phenotype of embryonic chicken cardiac myocytes. J. Biomed. Mater. Res., Part A 2010, 95A (4), 1261−1269. (22) Discher, D. E.; Janmey, P.; Wang, Y.-l. Tissue Cells Feel and Respond to the Stiffness of Their Substrate. Science 2005, 310 (5751), 1139−1143. (23) Trappmann, B.; Gautrot, J. E.; Connelly, J. T.; Strange, D. G. T.; Li, Y.; Oyen, M. L.; Cohen Stuart, M. A.; Boehm, H.; Li, B.; Vogel, V.; Spatz, J. P.; Watt, F. M.; Huck, W. T. S. Extracellular-matrix tethering regulates stem-cell fate. Nat. Mater. 2012, 11 (7), 642−649. (24) Bajaj, P.; Khang, D.; Webster, T. Control of spatial cell attachment on carbon nanofiber patterns on polycarbonate urethane. Int. J. Nanomed. 2006, 1 (3), 361−365. (25) Khetan, S.; Burdick, J. A. Patterning network structure to spatially control cellular remodeling and stem cell fate within 3dimensional hydrogels. Biomaterials 2010, 31 (32), 8228−8234. (26) Burdick, J. A.; Vunjak-Novakovic, G. Engineered microenvironments for controlled stem cell differentiation. Tissue Eng., Part A 2009, 15 (2), 205−19. (27) Tang, X.; Bajaj, P.; Bashir, R.; Saif, T. A. How far cardiac cells can see each other mechanically. Soft Matter 2011, 7 (13), 6151−6158. (28) Bajaj, P.; Marchwiany, D.; Duarte, C.; Bashir, R. Patterned three-dimensional encapsulation of embryonic stem cells using dielectrophoresis and stereolithography. Adv. Healthcare Mater. 2013, 2 (3), 450−8. (29) Mammoto, T.; Jiang, E.; Jiang, A.; Mammoto, A. Extracellular Matrix Structure and Tissue Stiffness Control Postnatal Lung Development through the Lipoprotein Receptor−Related Protein 5/ Tie2 Signaling System. Am. J. Respir. Cell Mol. Biol. 2013, 49 (6), 1009−1018. (30) Booth, A. J.; Hadley, R.; Cornett, A. M.; Dreffs, A. A.; Matthes, S. A.; Tsui, J. L.; Weiss, K.; Horowitz, J. C.; Fiore, V. F.; Barker, T. H.; Moore, B. B.; Martinez, F. J.; Niklason, L. E.; White, E. S. Acellular Normal and Fibrotic Human Lung Matrices as a Culture System for In Vitro Investigation. Am. J. Respir. Crit. Care Med. 2012, 186 (9), 866− 876. (31) Pelosi, P.; Rocco, P. Effects of mechanical ventilation on the extracellular matrix. Intensive Care Med. 2008, 34 (4), 631−639. (32) Faffe, D. S.; Zin, W. A. Lung Parenchymal Mechanics in Health and Disease. Physiol. Rev. 2009, 89 (3), 759−775. (33) Suki, B.; Bates, J. H. T. Extracellular matrix mechanics in lung parenchymal diseases. Respir. Physiol. Neurobiol. 2008, 163 (1−3), 33− 43. (34) Wynn, T. A. Integrating mechanisms of pulmonary fibrosis. J. Exp. Med. 2011, 208 (7), 1339−1350. (35) Kaminsky, D. A.; Irvin, C. G.; Lundblad, L.; Moriya, H. T.; Lang, S.; Allen, J.; Viola, T.; Lynn, M.; Bates, J. H. T. Oscillation mechanics of the human lung periphery in asthma. J. Appl. Physiol. 2004, 97 (5), 1849−1858.

‡

P.B. is currently at Primary Pharmacology Group, PDM-NCE Pfizer Inc., Groton, CT 06340. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This project was funded by the Defense Threat Reduction Agency (DTRA) program: Integration of Novel Technologies for Organ Development and Rapid Assessment of Medical Countermeasures (INTO-RAM), DTRA100271A5196. Los Alamos National Laboratory, an affirmative action equal opportunity employer, is operated by Los Alamos National Security, LLC, for the National Nuclear Security Administration of the U.S. Department of Energy under contract DEAC52- 06NA25396. The authors also thank Dr. John P. Wikswo at Vanderbilt University. The LA-UR number for the work is 14-24197.

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REFERENCES

(1) Boron, W. F. Organization of the Respiratory System, 1st ed.; Elsevier: Philadelphia, PA, 2005; p 1269. (2) Weibel, E. R., Geometric and Dimensional Airway Models of Conductive, Transitory and Respiratory Zones of the Human Lung. In Morphometry of the Human Lung; Weibel, E. R., Ed.; Academic Press: New York, 1963; Chapter XI, pp 136−142. DOI: 10.1016/B978-14832-0076-7.50016-8. (3) Douville, N. J. Alveolar Microfluidic Systems for the Study of Barrier Function, Cell Damage, and Migration at the Air−Blood Barrier. PhD Thesis, Univeristy of Michigan, Ann Arbor, MI, 2011. (4) Zarei, S.; Mirtar, A.; Andresen, B.; Salamon, P. Modeling the airflow in a lung with cystic fibrosis. J. Non-Equilib. Thermodyn. 2013, 38, 119. (5) Boron, W. F. Mechanics of Respiration. 1st ed.; Elsevier: Philadelphia, PA, 2005; p 1269. (6) Sherwood, L. Fundamentals of Human Physiology. 4th ed.; Cengage Learning: Independence, KY, 2011. (7) Maina, J. N. Fundamental structural aspects and features in the bioengineering of the gas exchangers: comparative perspectives. Adv. Anat., Embryol. Cell Biol. 2002, 163 (III-XII), 1−108. (8) McQualter, J. L.; Bertoncello, I. Concise review: Deconstructing the lung to reveal its regenerative potential. Stem Cells 2012, 30 (5), 811−6. (9) Shebani, E.; Shahana, S.; Janson, C.; Roomans, G. M. Attachment of columnar airway epithelial cells in asthma. Tissue Cell 2005, 37 (2), 145−152. (10) Woywodt, A.; Lefrak, S.; Matteson, E. Tainted eponyms in medicine: the “Clara” cell joins the list. Eur. Respir. J. 2010, 36 (4), 706−708. (11) Wagner, D. E.; Bonvillain, R. W.; Jensen, T.; Girard, E. D.; Bunnell, B. A.; Finck, C. M.; Hoffman, A. M.; Weiss, D. J. Can stem cells be used to generate new lungs? Ex vivo lung bioengineering with decellularized whole lung scaffolds. Respirology 2013, 18 (6), 895−911. (12) Hogan, B. L.; Barkauskas, C. E.; Chapman, H. A.; Epstein, J. A.; Jain, R.; Hsia, C. C.; Niklason, L.; Calle, E.; Le, A.; Randell, S. H.; Rock, J.; Snitow, M.; Krummel, M.; Stripp, B. R.; Vu, T.; White, E. S.; Whitsett, J. A.; Morrisey, E. E. Repair and regeneration of the respiratory system: complexity, plasticity, and mechanisms of lung stem cell function. Cell stem cell 2014, 15 (2), 123−38. (13) Ren, X.; Moser, P. T.; Gilpin, S. E.; Okamoto, T.; Wu, T.; Tapias, L. F.; Mercier, F. E.; Xiong, L.; Ghawi, R.; Scadden, D. T.; Mathisen, D. J.; Ott, H. C. Engineering pulmonary vasculature in decellularized rat and human lungs. Nat. Biotechnol. 2015, 33 (10), 1097−102. (14) Ward, H. E.; Nicholas, T. E. Alveolar type I and type II cells. Aust. N. Z. J. Med. 1984, 14 (5 Suppl 3), 731−734. L


Review

ACS Biomaterials Science & Engineering (36) Postma, D. S.; Timens, W. Remodeling in Asthma and Chronic Obstructive Pulmonary Disease. Proc. Am. Thorac. Soc. 2006, 3 (5), 434−439. (37) Brewster, C. E. P.; Howarth, P. H.; Djukanovic, R.; Wilson, J.; Holgate, S. T.; Roche, W. R. Myofibroblasts and Subepithelial Fibrosis in Bronchial Asthma. Am. J. Respir. Cell Mol. Biol. 1990, 3 (5), 507− 511. (38) Barnes, P. J. Chronic Obstructive Pulmonary Disease. N. Engl. J. Med. 2000, 343 (4), 269−280. (39) Gazdar, A. F.; Gao, B.; Minna, J. D. Lung cancer cell lines: Useless artifacts or invaluable tools for medical science? Lung cancer 2010, 68 (3), 309−18. (40) Gazdar, A. F.; Girard, L.; Lockwood, W. W.; Lam, W. L.; Minna, J. D. Lung cancer cell lines as tools for biomedical discovery and research. J. Natl. Cancer Inst. 2010, 102 (17), 1310−21. (41) Nalayanda, D. D.; Wang, Q.; Fulton, W. B.; Wang, T. H.; Abdullah, F. Engineering an artificial alveolar-capillary membrane: a novel continuously perfused model within microchannels. J. Pediatr. Surg. 2010, 45 (1), 45−51. (42) Kamotani, Y.; Bersano-Begey, T.; Kato, N.; Tung, Y. C.; Huh, D.; Song, J. W.; Takayama, S. Individually programmable cell stretching microwell arrays actuated by a Braille display. Biomaterials 2008, 29 (17), 2646−55. (43) Douville, N. J.; Zamankhan, P.; Tung, Y.-C.; Li, R.; Vaughan, B. L.; Tai, C.-F.; White, J.; Christensen, P. J.; Grotberg, J. B.; Takayama, S. Combination of fluid and solid mechanical stresses contribute to cell death and detachment in a microfluidic alveolar model. Lab Chip 2011, 11 (4), 609−619. (44) Deforet, M.; Parrini, M. C.; Petitjean, L.; Biondini, M.; Buguin, A.; Camonis, J.; Silberzan, P. Automated velocity mapping of migrating cell populations (AVeMap). Nat. Methods 2012, 9 (11), 1081−3. (45) Takai, E.; Tsukimoto, M.; Harada, H.; Sawada, K.; Moriyama, Y.; Kojima, S. Autocrine regulation of TGF-beta1-induced cell migration by exocytosis of ATP and activation of P2 receptors in human lung cancer cells. J. Cell Sci. 2012, 125 (21), 5051−5060. (46) Nalayanda, D. D.; Puleo, C.; Fulton, W. B.; Sharpe, L. M.; Wang, T. H.; Abdullah, F. An open-access microfluidic model for lungspecific functional studies at an air-liquid interface. Biomed. Microdevices 2009, 11 (5), 1081−9. (47) Ott, H. C.; Clippinger, B.; Conrad, C.; Schuetz, C.; Pomerantseva, I.; Ikonomou, L.; Kotton, D.; Vacanti, J. P. Regeneration and orthotopic transplantation of a bioartificial lung. Nat. Med. 2010, 16 (8), 927−33. (48) Petersen, T. H.; Calle, E. A.; Zhao, L.; Lee, E. J.; Gui, L.; Raredon, M. B.; Gavrilov, K.; Yi, T.; Zhuang, Z. W.; Breuer, C.; Herzog, E.; Niklason, L. E. Tissue-Engineered Lungs for in Vivo Implantation. Science 2010, 329 (5991), 538−541. (49) Wang, D.; Haviland, D. L.; Burns, A. R.; Zsigmond, E.; Wetsel, R. A. A pure population of lung alveolar epithelial type II cells derived from human embryonic stem cells. Proc. Natl. Acad. Sci. U. S. A. 2007, 104 (11), 4449−54. (50) Swain, R. J.; Kemp, S. J.; Goldstraw, P.; Tetley, T. D.; Stevens, M. M. Assessment of cell line models of primary human cells by Raman spectral phenotyping. Biophys. J. 2010, 98 (8), 1703−11. (51) Mason, R. J.; Williams, M. C. Phospholipid composition and ultrastructure of A549 cells and other cultured pulmonary epithelial cells of presumed type II cell origin. Biochim. Biophys. Acta, Lipids Lipid Metab. 1980, 617 (1), 36−50. (52) Nardone, L. L.; Andrews, S. B. Cell line A549 as a model of the type II pneumocyte: Phospholipid biosynthesis from native and organometallic precursors. Biochim. Biophys. Acta, Lipids Lipid Metab. 1979, 573 (2), 276−295. (53) Foster, K. A.; Oster, C. G.; Mayer, M. M.; Avery, M. L.; Audus, K. L. Characterization of the A549 Cell Line as a Type II Pulmonary Epithelial Cell Model for Drug Metabolism. Exp. Cell Res. 1998, 243 (2), 359−366. (54) Smith, B. Cell line A549: a model system for the study of alveolar type II cell function. Am. Rev. Respir. Dis. 1977, 115 (2), 285− 293.

(55) 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 1976, 17 (1), 62−70. (56) Balis, J. U.; Bumgarner, S. D.; Paciga, J. E.; Paterson, J. F.; Shelley, S. A. Synthesis of lung surfactant-associated glycoproteins by A549 cells: description of an in vitro model for human type II cell dysfunction. Exp. Lung Res. 1984, 6 (3−4), 197−213. (57) Hermanns, M. I.; Unger, R. E.; Kehe, K.; Peters, K.; Kirkpatrick, C. J. Lung epithelial cell lines in coculture with human pulmonary microvascular endothelial cells: development of an alveolo-capillary barrier in vitro. Lab. Invest. 2004, 84 (6), 736−52. (58) Fulcher, M. L.; Gabriel, S.; Burns, K. A.; Yankaskas, J. R.; Randell, S. H. Well-differentiated human airway epithelial cell cultures. Methods Mol. Med. 2005, 107, 183−206. (59) US-FDA CFR, Code of Federal Regulations Title 21; http://www. accessdata.fda.gov/scripts/cdrh/cfdocs/cfcfr /CFRSearch. cfm?CFRPart=1271&showFR=1&subpartNode=21:8.0.1.5.55.3 (accessed March 16, 2014). (60) Lin, H.; Li, H.; Cho, H. J.; Bian, S.; Roh, H. J.; Lee, M. K.; Kim, J. S.; Chung, S. J.; Shim, C. K.; Kim, D. D. Air-liquid interface (ALI) culture of human bronchial epithelial cell monolayers as an in vitro model for airway drug transport studies. J. Pharm. Sci. 2007, 96 (2), 341−50. (61) Avadhanula, V.; Rodriguez, C. A.; Devincenzo, J. P.; Wang, Y.; Webby, R. J.; Ulett, G. C.; Adderson, E. E. Respiratory viruses augment the adhesion of bacterial pathogens to respiratory epithelium in a viral species- and cell type-dependent manner. J. Virol. 2006, 80 (4), 1629−1636. (62) Aarbiou, J.; Copreni, E.; Buijs-Offerman, R. M.; van der Wegen, P.; Castellani, S.; Carbone, A.; Tilesi, F.; Fradiani, P.; Hiemstra, P. S.; Yueksekdag, G.; Diana, A.; Rosenecker, J.; Ascenzioni, F.; Conese, M.; Scholte, B. J. Lentiviral small hairpin RNA delivery reduces apical sodium channel activity in differentiated human airway epithelial cells. journal of gene medicine 2012, 14 (12), 733−45. (63) Ng, C. T.; Li, J. J.; Gurung, R. L.; Hande, M. P.; Ong, C. N.; Bay, B. H.; Yung, L. Y. Toxicological profile of small airway epithelial cells exposed to gold nanoparticles. Exp. Biol. Med. 2013, 238 (12), 1355−61. (64) Pirela, S. V.; Miousse, I. R.; Lu, X.; Castranova, V.; Thomas, T.; Qian, Y.; Bello, D.; Kobzik, L.; Koturbash, I.; Demokritou, P. Effects of Laser Printer-Emitted Engineered Nanoparticles on Cytotoxicity, Chemokine Expression, Reactive Oxygen Species, DNA Methylation, and DNA Damage: A Comprehensive Analysis in Human Small Airway Epithelial Cells, Macrophages, and Lymphoblasts. Environ. Health Perspect. 2016, DOI: 10.1289/ehp.1409582. (65) Tang, M.; Li, Q.; Xiao, L.; Li, Y.; Jensen, J. L.; Liou, T. G.; Zhou, A. Toxicity effects of short term diesel exhaust particles exposure to human small airway epithelial cells (SAECs) and human lung carcinoma epithelial cells (A549). Toxicol. Lett. 2012, 215 (3), 181− 92. (66) Lee, J. W.; Fang, X.; Krasnodembskaya, A.; Howard, J. P.; Matthay, M. A. Concise review: Mesenchymal stem cells for acute lung injury: role of paracrine soluble factors. Stem Cells 2011, 29 (6), 913− 9. (67) Hayes, M.; Curley, G.; Ansari, B.; Laffey, J. G. Clinical review: Stem cell therapies for acute lung injury/acute respiratory distress syndrome - hope or hype? Critical care 2012, 16 (2), 205. (68) Bonvillain, R. W.; Danchuk, S.; Sullivan, D. E.; Betancourt, A. M.; Semon, J. A.; Eagle, M. E.; Mayeux, J. P.; Gregory, A. N.; Wang, G.; Townley, I. K.; Borg, Z. D.; Weiss, D. J.; Bunnell, B. A. A nonhuman primate model of lung regeneration: detergent-mediated decellularization and initial in vitro recellularization with mesenchymal stem cells. Tissue Eng., Part A 2012, 18 (23−24), 2437−2452. (69) Wagner, D. E.; Bonenfant, N. R.; Parsons, C. S.; Sokocevic, D.; Brooks, E. M.; Borg, Z. D.; Lathrop, M. J.; Wallis, J. D.; Daly, A. B.; Lam, Y. W.; Deng, B.; DeSarno, M. J.; Ashikaga, T.; Loi, R.; Weiss, D. J. Comparative decellularization and recellularization of normal versus emphysematous human lungs. Biomaterials 2014, 35 (10), 3281−3297. M


Review


(91) Geerts, H. Of Mice and Men: Bridging the Translational Disconnect in CNS Drug Discovery. CNS Drugs 2009, 23 (11), 915− 926. (92) Esch, M. B.; King, T. L.; Shuler, M. L. The Role of Body-on-aChip Devices in Drug and Toxicity Studies. Annu. Rev. Biomed. Eng. 2011, 13 (1), 55−72. (93) van de Stolpe, A.; den Toonder, J. Workshop meeting report Organs-on-Chips: human disease models. Lab Chip 2013, 13 (18), 3449−3470. (94) Tesfaigzi, Y. Roles of apoptosis in airway epithelia. Am. J. Respir. Cell Mol. Biol. 2006, 34 (5), 537−547. (95) Hauber, H. P.; Foley, S. C.; Hamid, Q. Mucin overproduction in chronic inflammatory lung disease. Can. Respir. J. 2006, 13 (6), 327− 335. (96) Viravaidya, K.; Sin, A.; Shuler, M. L. Development of a Microscale Cell Culture Analog To Probe Naphthalene Toxicity. Biotechnol. Prog. 2004, 20 (1), 316−323. (97) Sung, J. H.; Shuler, M. L. A micro cell culture analog ([small micro]CCA) with 3-D hydrogel culture of multiple cell lines to assess metabolism-dependent cytotoxicity of anti-cancer drugs. Lab Chip 2009, 9 (10), 1385−1394. (98) Moraes, C.; Labuz, J. M.; Leung, B. M.; Inoue, M.; Chun, T.-H.; Takayama, S. On being the right size: scaling effects in designing a human-on-a-chip. Integrative Biology 2013, 5 (9), 1149−1161. (99) West, G. B.; Brown, J. H.; Enquist, B. J. A General Model for the Origin of Allometric Scaling Laws in Biology. Science 1997, 276 (5309), 122−126. (100) Wikswo, J. P.; Curtis, E. L.; Eagleton, Z. E.; Evans, B. C.; Kole, A.; Hofmeister, L. H.; Matloff, W. J. Scaling and systems biology for integrating multiple organs-on-a-chip. Lab Chip 2013, 13 (18), 3496− 3511. (101) Huang, Y.; Williams, J. C.; Johnson, S. M. Brain slice on a chip: opportunities and challenges of applying microfluidic technology to intact tissues. Lab Chip 2012, 12 (12), 2103−2117. (102) Maher, M.; Pine, J.; Wright, J.; Tai, Y.-C. The neurochip: a new multielectrode device for stimulating and recording from cultured neurons. J. Neurosci. Methods 1999, 87 (1), 45−56. (103) Jang, K.-J.; Mehr, A. P.; Hamilton, G. A.; McPartlin, L. A.; Chung, S.; Suh, K.-Y.; Ingber, D. E. Human kidney proximal tubuleon-a-chip for drug transport and nephrotoxicity assessment. Integrative Biology 2013, 5 (9), 1119−1129. (104) Jang, K.-J.; Suh, K.-Y. A multi-layer microfluidic device for efficient culture and analysis of renal tubular cells. Lab Chip 2010, 10 (1), 36−42. (105) Kim, H. J.; Huh, D.; Hamilton, G.; Ingber, D. E. Human guton-a-chip inhabited by microbial flora that experiences intestinal peristalsis-like motions and flow. Lab Chip 2012, 12 (12), 2165−2174. (106) Grosberg, A.; Alford, P. W.; McCain, M. L.; Parker, K. K. Ensembles of engineered cardiac tissues for physiological and pharmacological study: Heart on a chip. Lab Chip 2011, 11 (24), 4165−4173. (107) Goral, V. N.; Hsieh, Y.-C.; Petzold, O. N.; Clark, J. S.; Yuen, P. K.; Faris, R. A. Perfusion-based microfluidic device for threedimensional dynamic primary human hepatocyte cell culture in the absence of biological or synthetic matrices or coagulants. Lab Chip 2010, 10 (24), 3380−3386. (108) Ho, C.-T.; Lin, R.-Z.; Chen, R.-J.; Chin, C.-K.; Gong, S.-E.; Chang, H.-Y.; Peng, H.-L.; Hsu, L.; Yew, T.-R.; Chang, S.-F.; Liu, C.H. Liver-cell patterning Lab Chip: mimicking the morphology of liver lobule tissue. Lab Chip 2013, 13 (18), 3578−3587. (109) Lee, S.-A.; No, D. Y.; Kang, E.; Ju, J.; Kim, D.-S.; Lee, S.-H. Spheroid-based three-dimensional liver-on-a-chip to investigate hepatocyte-hepatic stellate cell interactions and flow effects. Lab Chip 2013, 13 (18), 3529−3537. (110) Huh, D.; Leslie, D. C.; Matthews, B. D.; Fraser, J. P.; Jurek, S.; Hamilton, G. A.; Thorneloe, K. S.; McAlexander, M. A.; Ingber, D. E. A Human Disease Model of Drug Toxicity−Induced Pulmonary Edema in a Lung-on-a-Chip Microdevice. Sci. Transl. Med. 2012, 4 (159), 159ra147.

(70) Huang, S. X. L.; Islam, M. N.; O’Neill, J.; Hu, Z.; Yang, Y.-G.; Chen, Y.-W.; Mumau, M.; Green, M. D.; Vunjak-Novakovic, G.; Bhattacharya, J.; Snoeck, H.-W. Efficient generation of lung and airway epithelial cells from human pluripotent stem cells. Nat. Biotechnol. 2014, 32 (1), 84−91. (71) Hattori, K.; Sugiura, S.; Kanamori, T. Microenvironment array chip for cell culture environment screening. Lab Chip 2011, 11 (2), 212−214. (72) El-Mounayri, O.; Mihic, A.; Shikatani, E. A.; Gagliardi, M.; Steinbach, S. K.; Dubois, N.; Dacosta, R.; Li, R. K.; Keller, G.; Husain, M. Serum-free differentiation of functional human coronary-like vascular smooth muscle cells from embryonic stem cells. Cardiovasc. Res. 2013, 98 (1), 125−35. (73) Fraslon, C.; Rolland, G.; Bourbon, J. R.; Rieutort, M.; Valenza, C. Culture of fetal alveolar epithelial type II cells in serum-free medium. In Vitro Cell. Dev. Biol. 1991, 27A (11), 843−852. (74) Winkler, M. E.; Mauritz, C.; Groos, S.; Kispert, A.; Menke, S.; Hoffmann, A.; Gruh, I.; Schwanke, K.; Haverich, A.; Martin, U. Serumfree differentiation of murine embryonic stem cells into alveolar type II epithelial cells. Cloning Stem Cells 2008, 10 (1), 49−64. (75) Suki, B.; Ito, S.; Stamenović, D.; Lutchen, K. R.; Ingenito, E. P. Biomechanics of the lung parenchyma: critical roles of collagen and mechanical forces. J. Appl. Physiol. 2005, 98 (5), 1892−1899. (76) Huh, D.; Kim, H. J.; Fraser, J. P.; Shea, D. E.; Khan, M.; Bahinski, A.; Hamilton, G. A.; Ingber, D. E. Microfabrication of human organs-on-chips. Nat. Protoc. 2013, 8 (11), 2135−2157. (77) Huh, D.; Hamilton, G. A.; Ingber, D. E. From 3D cell culture to organs-on-chips. Trends Cell Biol. 2011, 21 (12), 745−754. (78) Jang, K. J.; Cho, H. S.; Kang do, H.; Bae, W. G.; Kwon, T. H.; Suh, K. Y. Fluid-shear-stress-induced translocation of aquaporin-2 and reorganization of actin cytoskeleton in renal tubular epithelial cells. Integr Biol. (Camb) 2011, 3 (2), 134−141. (79) Sciancalepore, A. G.; Sallustio, F.; Girardo, S.; Gioia Passione, L.; Camposeo, A.; Mele, E.; Di Lorenzo, M.; Costantino, V.; Schena, F. P.; Pisignano, D. A bioartificial renal tubule device embedding human renal stem/progenitor cells. PLoS One 2014, 9 (1), e87496. (80) Hoffmann, S. A.; Muller-Vieira, U.; Biemel, K.; Knobeloch, D.; Heydel, S.; Lubberstedt, M.; Nussler, A. K.; Andersson, T. B.; Gerlach, J. C.; Zeilinger, K. Analysis of drug metabolism activities in a miniaturized liver cell bioreactor for use in pharmacological studies. Biotechnol. Bioeng. 2012, 109 (12), 3172−81. (81) Kim, H. J.; Ingber, D. E. Gut-on-a-Chip microenvironment induces human intestinal cells to undergo villus differentiation. Integrative Biology 2013, 5 (9), 1130−1140. (82) Huh, D.; Torisawa, Y.-s.; Hamilton, G. A.; Kim, H. J.; Ingber, D. E. Microengineered physiological biomimicry: Organs-on-Chips. Lab Chip 2012, 12 (12), 2156−2164. (83) Huh, D.; Matthews, B. D.; Mammoto, A.; Montoya-Zavala, M.; Hsin, H. Y.; Ingber, D. E. Reconstituting Organ-Level Lung Functions on a Chip. Science 2010, 328 (5986), 1662−1668. (84) Agarwal, A.; Goss, J. A.; Cho, A.; McCain, M. L.; Parker, K. K. Microfluidic heart on a chip for higher throughput pharmacological studies. Lab Chip 2013, 13 (18), 3599−3608. (85) Collins, F. S. Reengineering Translational Science: The Time Is Right. Sci. Transl. Med. 2011, 3 (90), 90cm17. (86) Bolten, B. M.; DeGregorio, T. Trends in development cycles. Nat. Rev. Drug Discovery 2002, 1 (5), 335−336. (87) Paul, S. M.; Mytelka, D. S.; Dunwiddie, C. T.; Persinger, C. C.; Munos, B. H.; Lindborg, S. R.; Schacht, A. L. How to improve R&D productivity: the pharmaceutical industry’s grand challenge. Nat. Rev. Drug Discovery 2010, 9 (3), 203−214. (88) Munos, B. Lessons from 60 years of pharmaceutical innovation. Nat. Rev. Drug Discovery 2009, 8 (12), 959−968. (89) Scannell, J. W.; Blanckley, A.; Boldon, H.; Warrington, B. Diagnosing the decline in pharmaceutical R&D efficiency. Nat. Rev. Drug Discovery 2012, 11 (3), 191−200. (90) de Jong, M.; Maina, T. Of Mice and Humans: Are They the Same?Implications in Cancer Translational Research. J. Nucl. Med. 2010, 51 (4), 501−504. N


Review

ACS Biomaterials Science & Engineering (111) Thorneloe, K. S.; Cheung, M.; Bao, W.; Alsaid, H.; Lenhard, S.; Jian, M. Y.; Costell, M.; Maniscalco-Hauk, K.; Krawiec, J. A.; Olzinski, A.; Gordon, E.; Lozinskaya, I.; Elefante, L.; Qin, P.; Matasic, D. S.; James, C.; Tunstead, J.; Donovan, B.; Kallal, L.; Waszkiewicz, A.; Vaidya, K.; Davenport, E. A.; Larkin, J.; Burgert, M.; Casillas, L. N.; Marquis, R. W.; Ye, G.; Eidam, H. S.; Goodman, K. B.; Toomey, J. R.; Roethke, T. J.; Jucker, B. M.; Schnackenberg, C. G.; Townsley, M. I.; Lepore, J. J.; Willette, R. N. An orally active TRPV4 channel blocker prevents and resolves pulmonary edema induced by heart failure. Sci. Transl. Med. 2012, 4 (159), 159ra148. (112) Huh, D.; Fujioka, H.; Tung, Y.-C.; Futai, N.; Paine, R.; Grotberg, J. B.; Takayama, S. Acoustically detectable cellular-level lung injury induced by fluid mechanical stresses in microfluidic airway systems. Proc. Natl. Acad. Sci. U. S. A. 2007, 104 (48), 18886−18891. (113) Slutsky, A. S.; Ranieri, V. M. Ventilator-Induced Lung Injury. N. Engl. J. Med. 2013, 369 (22), 2126−2136. (114) Xu, Z.; Gao, Y.; Hao, Y.; Li, E.; Wang, Y.; Zhang, J.; Wang, W.; Gao, Z.; Wang, Q. Application of a microfluidic chip-based 3D coculture to test drug sensitivity for individualized treatment of lung cancer. Biomaterials 2013, 34 (16), 4109−4117. (115) Sellgren, K. L.; Butala, E. J.; Gilmour, B. P.; Randell, S. H.; Grego, S. A biomimetic multicellular model of the airways using primary human cells. Lab Chip 2014, 14, 3349. (116) Stucki, A. O.; Stucki, J. D.; Hall, S. R.; Felder, M.; Mermoud, Y.; Schmid, R. A.; Geiser, T.; Guenat, O. T. A lung-on-a-chip array with an integrated bio-inspired respiration mechanism. Lab Chip 2015, 15 (5), 1302−1310. (117) Whitesides, G. M.; Ostuni, E.; Takayama, S.; Jiang, X.; Ingber, D. E. SOFT LITHOGRAPHY IN BIOLOGY AND BIOCHEMISTRY. Annu. Rev. Biomed. Eng. 2001, 3 (1), 335−373. (118) Duffy, D. C.; McDonald, J. C.; Schueller, O. J. A.; Whitesides, G. M. Rapid Prototyping of Microfluidic Systems in Poly(dimethylsiloxane). Anal. Chem. 1998, 70 (23), 4974−4984. (119) Tsao, C.-W.; DeVoe, D. Bonding of thermoplastic polymer microfluidics. Microfluid. Nanofluid. 2009, 6 (1), 1−16. (120) Kuncova-Kallio, J.; Kallio, P. J. PDMS and Its Suitability for Analytical Microfluidic Devices. In 28th Annual International Conference on Engineering in Medicine and Biology Society; New York, Aug 30− Sept 3, 2006; IEEE: Piscataway, NJ, 2006; pp 2486−2489, DOI: 10.1109/IEMBS.2006.260465. (121) Merkel, T. C.; Bondar, V. I.; Nagai, K.; Freeman, B. D.; Pinnau, I. Gas sorption, diffusion, and permeation in poly(dimethylsiloxane). J. Polym. Sci., Part B: Polym. Phys. 2000, 38 (3), 415−434. (122) Fujii, T. PDMS-based microfluidic devices for biomedical applications. Microelectron. Eng. 2002, 61−62, 907−914. (123) Wang, J.; Douville, N.; Takayama, S.; ElSayed, M. Quantitative Analysis of Molecular Absorption into PDMS Microfluidic Channels. Ann. Biomed. Eng. 2012, 40 (9), 1862−1873. (124) Toepke, M. W.; Beebe, D. J. PDMS absorption of small molecules and consequences in microfluidic applications. Lab Chip 2006, 6 (12), 1484−6. (125) Gomez-Sjoberg, R.; Leyrat, A. A.; Houseman, B. T.; Shokat, K.; Quake, S. R. Biocompatibility and Reduced Drug Absorption of Sol− Gel-Treated Poly(dimethyl siloxane) for Microfluidic Cell Culture Applications. Anal. Chem. 2010, 82 (21), 8954−8960. (126) Zhou, J.; Ellis, A. V.; Voelcker, N. H. Recent developments in PDMS surface modification for microfluidic devices. Electrophoresis 2010, 31 (1), 2−16. (127) Wu, D.; Zhao, B.; Dai, Z.; Qin, J.; Lin, B. Grafting epoxymodified hydrophilic polymers onto poly(dimethylsiloxane) microfluidic chip to resist nonspecific protein adsorption. Lab Chip 2006, 6 (7), 942−7. (128) Thakar, R.; Zakeri, R.; Morris, C. A.; Baker, L. A. Rapid fabrication of nanoporous membrane arrays and single-pore membranes from parylene C. Anal. Methods 2012, 4 (12), 4353−4359. (129) Jinno, S.; Moeller, H.-C.; Chen, C.-L.; Rajalingam, B.; Chung, B. G.; Dokmeci, M. R.; Khademhosseini, A. Microfabricated multilayer parylene-C stencils for the generation of patterned dynamic cocultures. J. Biomed. Mater. Res., Part A 2008, 86A (1), 278−288.

(130) Wright, D.; Rajalingam, B.; Selvarasah, S.; Dokmeci, M. R.; Khademhosseini, A. Generation of static and dynamic patterned cocultures using microfabricated parylene-C stencils. Lab Chip 2007, 7 (10), 1272−1279. (131) Lu, B.; Zhu, D.; Hinton, D.; Humayun, M.; Tai, Y.-C. Meshsupported submicron parylene-C membranes for culturing retinal pigment epithelial cells. Biomed. Microdevices 2012, 14 (4), 659−667. (132) Gutierrez, C. A.; Meng, E. Liquid Encapsulation in Parylene Microstructures Using Integrated Annular-Plate Stiction Valves. Micromachines 2011, 2 (3), 356−368. (133) Kasper, J.; Hermanns, M. I.; Bantz, C.; Maskos, M.; Stauber, R.; Pohl, C.; Unger, R. E.; Kirkpatrick, J. C. Inflammatory and cytotoxic responses of an alveolar-capillary coculture model to silica nanoparticles: comparison with conventional monocultures. Part. Fibre Toxicol. 2011, 8 (1), 6. (134) Engler, A. J.; Sen, S.; Sweeney, H. L.; Discher, D. E. Matrix Elasticity Directs Stem Cell Lineage Specification. Cell 2006, 126 (4), 677−689. (135) Kilian, K. A.; Bugarija, B.; Lahn, B. T.; Mrksich, M. Geometric cues for directing the differentiation of mesenchymal stem cells. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 4872. (136) Nelson, C. M.; Jean, R. P.; Tan, J. L.; Liu, W. F.; Sniadecki, N. J.; Spector, A. A.; Chen, C. S. Emergent patterns of growth controlled by multicellular form and mechanics. Proc. Natl. Acad. Sci. U. S. A. 2005, 102 (33), 11594−11599. (137) McBeath, R.; Pirone, D. M.; Nelson, C. M.; Bhadriraju, K.; Chen, C. S. Cell Shape, Cytoskeletal Tension, and RhoA Regulate Stem Cell Lineage Commitment. Dev. Cell 2004, 6 (4), 483−495. (138) Berthiaume, F.; Moghe, P. V.; Toner, M.; Yarmush, M. L. Effect of extracellular matrix topology on cell structure, function, and physiological responsiveness: hepatocytes cultured in a sandwich configuration. FASEB J. 1996, 10 (13), 1471−1484. (139) Ross, A. M.; Jiang, Z.; Bastmeyer, M.; Lahann, J. Physical Aspects of Cell Culture Substrates: Topography, Roughness, and Elasticity. Small 2012, 8 (3), 336−355. (140) Charest, J. L.; García, A. J.; King, W. P. Myoblast alignment and differentiation on cell culture substrates with microscale topography and model chemistries. Biomaterials 2007, 28 (13), 2202−2210. (141) Li, H.; Wijekoon, A.; Leipzig, N. D. 3D Differentiation of Neural Stem Cells in Macroporous Photopolymerizable Hydrogel Scaffolds. PLoS One 2012, 7 (11), e48824. (142) Chiu, Y. C.; Cheng, M. H.; Engel, H.; Kao, S. W.; Larson, J. C.; Gupta, S.; Brey, E. M. The role of pore size on vascularization and tissue remodeling in PEG hydrogels. Biomaterials 2011, 32 (26), 6045−51. (143) Borok, Z.; Crandall, E. D. More life for a “terminal” cell. Am. J. Physiol: Lung Cell. Mol. Physiol. 2009, 297, L1042−L1044. (144) Bajaj, P.; Schweller, R. M.; Khademhosseini, A.; West, J. L.; Bashir, R. 3D Biofabrication Strategies for Tissue Engineering and Regenerative Medicine. Annu. Rev. Biomed. Eng. 2014, 16 (1), 247− 276. (145) Matsui, H.; Grubb, B. R.; Tarran, R.; Randell, S. H.; Gatzy, J. T.; Davis, C. W.; Boucher, R. C. Evidence for Periciliary Liquid Layer Depletion, Not Abnormal Ion Composition, in the Pathogenesis of Cystic Fibrosis Airways Disease. Cell 1998, 95 (7), 1005−1015. (146) Matsui, H.; Randell, S. H.; Peretti, S. W.; Davis, C. W.; Boucher, R. C. Coordinated clearance of periciliary liquid and mucus from airway surfaces. J. Clin. Invest. 1998, 102 (6), 1125−1131. (147) Grubb, B. R.; Schiretz, F. R.; Boucher, R. C. Volume transport across tracheal and bronchial airway epithelia in a tubular culture system. Am. J. Physiol.: Cell Physiol. 1997, 273 (1), C21−C29. (148) Ryu, J. H.; Myers, J. L.; Swensen, S. J. Bronchiolar Disorders. Am. J. Respir. Crit. Care Med. 2003, 168 (11), 1277−1292. (149) Guo, X.; Gonzalez, M.; Stancescu, M.; Vandenburgh, H. H.; Hickman, J. J. Neuromuscular junction formation between human stem cell-derived motoneurons and human skeletal muscle in a defined system. Biomaterials 2011, 32 (36), 9602−9611. O


Review

ACS Biomaterials Science & Engineering (150) Smith, A.; Long, C.; Berry, B.; McAleer, C.; Stancescu, M.; Molnar, P.; Miller, P.; Esch, M.; Prot, J.-M.; Hickman, J.; Shuler, M. Microphysiological systems and low-cost microfluidic platform with analytics. Stem Cell Res. Ther. 2013, 4 (Suppl 1), S9. (151) Mosadegh, B.; Mazzeo, A. D.; Shepherd, R. F.; Morin, S. A.; Gupta, U.; Sani, I. Z.; Lai, D.; Takayama, S.; Whitesides, G. M. Control of soft machines using actuators operated by a Braille display. Lab Chip 2014, 14 (1), 189−199. (152) Huh, D.; Kim, H. J.; Fraser, J. P.; Shea, D. E.; Khan, M.; Bahinski, A.; Hamilton, G. A.; Ingber, D. E. Microfabrication of human organs-on-chips. Nat. Protoc. 2013, 8 (11), 2135−57. (153) Benam, K. H.; Villenave, R.; Lucchesi, C.; Varone, A.; Hubeau, C.; Lee, H. H.; Alves, S. E.; Salmon, M.; Ferrante, T. C.; Weaver, J. C.; Bahinski, A.; Hamilton, G. A.; Ingber, D. E. Small airway-on-a-chip enables analysis of human lung inflammation and drug responses in vitro. Nat. Methods 2016, 13 (2), 151−157. (154) Huh, D.; Fujioka, H.; Tung, Y. C.; Futai, N.; Paine, R., 3rd; Grotberg, J. B.; Takayama, S. Acoustically detectable cellular-level lung injury induced by fluid mechanical stresses in microfluidic airway systems. Proc. Natl. Acad. Sci. U. S. A. 2007, 104 (48), 18886−91. (155) Tesei, A.; Zoli, W.; Arienti, C.; Storci, G.; Granato, A. M.; Pasquinelli, G.; Valente, S.; Orrico, C.; Rosetti, M.; Vannini, I.; Dubini, A.; Dell’Amore, D.; Amadori, D.; Bonafe, M. Isolation of stem/ progenitor cells from normal lung tissue of adult humans. Cell Proliferation 2009, 42 (3), 298−308. (156) Huh, D.; Matthews, B. D.; Mammoto, A.; Montoya-Zavala, M.; Hsin, H. Y.; Ingber, D. E. Reconstituting organ-level lung functions on a chip. Science 2010, 328 (5986), 1662−8. (157) Douville, N. J.; Zamankhan, P.; Tung, Y. C.; Li, R.; Vaughan, B. L.; Tai, C. F.; White, J.; Christensen, P. J.; Grotberg, J. B.; Takayama, S. Combination of fluid and solid mechanical stresses contribute to cell death and detachment in a microfluidic alveolar model. Lab Chip 2011, 11 (4), 609−19. (158) Punde, T. H.; Wu, W. H.; Lien, P. C.; Chang, Y. L.; Kuo, P. H.; Chang, M. D.; Lee, K. Y.; Huang, C. D.; Kuo, H. P.; Chan, Y. F.; Shih, P. C.; Liu, C. H. A biologically inspired lung-on-a-chip device for the study of protein-induced lung inflammation. Integr Biol. (Camb) 2015, 7 (2), 162−169.

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