Annals of Biomedical Engineering, Vol. 40, No. 6, June 2012 (Ó 2012) pp. 1289–1300 DOI: 10.1007/s10439-011-0491-2

Microtechnology for Mimicking In Vivo Tissue Environment JONG HWAN SUNG1 and MICHAEL L. SHULER2 1

Chemical Engineering, Hongik University, Seoul, Korea; and 2Biomedical Engineering, Cornell University, 115 Weill Hall, Ithaca, NY 14850, USA (Received 18 August 2011; accepted 14 December 2011; published online 4 January 2012) Associate Editor Tingrui Pan oversaw the review of this article.

In the natural tissue environment, cells are surrounded by 3-dimensional (3-D) organization of supporting matrix and neighboring cells, and a gradient of chemical and mechanical signals. The presence of blood flow and mechanical movement provides a dynamic environment and interactions between multiple organs often affect cell behavior. On the other hand, in vitro environment typically provides a static environment, usually with a single cell type, cultured on a 2-dimensional (2-D) monolayer. Microtechnology, which originated from the semiconductor industry, allows fabrication of structures in micrometer scale and integration of those structures on a small chip. During the past few decades, it has been realized that the microtechnology could be applied to analytical chemistry, which gave rise to the new area of microfluidics. Initially, the main advantages of microfluidics were thought to be reduced sample consumption, faster response time, increased resolution, and the possibility of high-throughput implementation.12 However, the true merit of the microscale systems comes from their spatiotemporal control of substances in micrometer scale.24 This ability enables accurate reconstruction of the biological context, which has a typical length scale of micrometers. Furthermore, microfluidics, which handles small volumes of liquid, can be used to mimic the dynamic environment of biological systems, such as blood circulation and gas transport. In this review, we summarize recent efforts in microtechnology aimed at reconstructing accurate analogs of in vivo environment. The main focus of this article will be on how microtechnology can be used to mimic and replicate human tissue environment and more importantly, whole-body response to drugs. By mimicking the biological context more accurately, cells are placed in a more physiologically realistic environment than traditional 2-D in vitro systems, and are likely to behave in more authentic manner. Microfabricated

Abstract—Microtechnology provides a new approach for reproducing the in vivo environment in vitro. Mimicking the microenvironment of the natural tissues allows cultured cells to behave in a more authentic manner, and gives researchers more realistic platforms to study biological systems. In this review article, we discuss the physiochemical aspects of in vivo cellular microenvironment, and relevant technologies that can be used to mimic those aspects. Secondly we identify the core methods used in microtechnology for biomedical applications. Finally we examine the recent application areas of microtechnology, with a focus on reproducing the functions of specific organs, or whole-body response such as homeostasis or metabolism-dependent toxicity of drugs. These new technologies enable researchers to ask and answer questions in a manner that has not been possible with conventional, macroscale technologies. Keywords—Microtechnology, In vitro systems, Multi-organ interactions, Pharmacokinetics, Microfluidics.

INTRODUCTION In vitro systems are isolated from their original biological context to allow controlled experimentation and analysis. In vitro systems are widely used in research for drug development, regenerative medicine, and tissue engineering, and to gain a better understanding of tissue and cell biology. Among the in vitro systems, cell culture models are well established technique, where cells are isolated from specific tissues, and cultured in a controlled human-designed environment. Recent studies reveal that the differences in the microenvironment provided by cell culture models and that in the in vivo tissues are significant and can cause deviations in cell response and behavior.10 The differences between in vivo and in vitro models are diverse. Address correspondence to Michael L. Shuler, Biomedical Engineering, Cornell University, 115 Weill Hall, Ithaca, NY 14850, USA. Electronic mail: [email protected]

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systems can mimic nanoscale surface patterns, release of chemical signals, and 3-D tissue organization. Combination of such biomimetic organs can be used to predict whole-body response to chemicals and drugs. Here, we first summarize different aspects of in vivo tissue environment and explain how microtechnology can be used to replicate such aspects. Then the methods of microtechnology applied to recreating 3-D tissue constructs are described, with examples of applications, and finally we conclude with key remaining challenges in developing microscale systems.

CELLULAR MICROENVIRONMENT AND RELEVANT TECHNOLOGIES In a natural tissue environment, cells are surrounded by various chemical, mechanical and physical cues (Fig. 1). These cues provide cells with a biological context, which acts to regulate the behavior of cells. More importantly, these factors can change in a timedependent manner, constituting a dynamic environment. Here we list the factors that constitute the in vivo cellular environment, and summarize relevant microtechnologies aimed at mimicking those factors. Chemical Gradient Gradients play a key role in determining the function and the fate of cells in vivo. Processes that are

FIGURE 1. Important factors of cellular microenvironment.

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affected by gradients include cell migration, angiogenesis, development and wound healing. Chemical gradients are typically formed as a result of diffusion, while mechanical gradients in substrate stiffness or ligands are formed by the heterogeneity of surrounding matrix. Microtechnology is an ideal tool for recreating these gradients, with its microscale control over transport. Typically, diffusion in the laminar flow is utilized to create desired concentration gradients of soluble factors or oxygen. The gradients can be linear, logarithmic, or arbitrary desired patterns.35,40 In the simplest form, fluidic streams from two separate sources can be merged together, and the laminar flow allows diffusive mixing between the two streams. Using such a method, Ismagilov and co-workers42 exposed a developing embryo of Drosophila to a step change in temperature to observe the response of the embryo to the artificial perturbation in the development rate. While initial approaches were focused on creating diverse forms of gradients in 2-D, it is obvious that a gradient across 3-D space mimic in vivo tissue environment better. Creation of a gradient in 3-D hydrogel matrix is achieved by placing the gel matrix between a source and a sink, followed by diffusive transport of molecules.58,79 The basic principles of gradient formation in 3-D is essentially the same as 2-D, but the formation process also depends on the permeability of the hydrogel matrix. Recent efforts in creating and using artificial concentration gradients (soluble factors or surface proteins) have been reviewed.28,32

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Gradients are not only applicable to chemicals such as growth factors and hormones, but also to the mechanical property of ECM or surface-bound ligands. Depending on the factors, motility of cells due to these factors are classified as chemotaxis (chemicals), durotaxis (substrate stiffness), and haptotaxis (surface-bound ligands). The effect of a gradient in substrate stiffness on bone marrow stromal cell differentiation was examined using commercially available hydrogel.61 It was observed that osteogenic differentiation decreased on substrates of lower stiffness and lower ligand density. Novel methods for forming a gradient in substrate stiffness have been developed to study the durotactic migration of cells. A microfluidic device was designed to form a gradient of stiffness in collagen gel, and neurites grew significantly longer down the gradient than up the gradient.67 In another study, mesenchymal stem cell differentiation in stiffness gradient was tested.77 Directed migration of cells up the stiffness gradients was observed. 3-D Architecture In tissue environment, cells are surrounded by 3-D organization of extracellular matrix (ECM) and neighboring cells, which interact to provide complex chemical and mechanical signals. Anchoring of cells to surrounding matrix via surface integrins is thought be responsible for not only the physical attachment, but also for transducing various chemical and mechanical signals from the outer environment.25 Recent studies have confirmed that cells cultured in 3-D environment indeed behave differently from those cultured in conventional 2-D culture.10,82 Bissel is one of the pioneers who elucidated the role of 3-D cell culture and demonstrated the basis for many of the methodologies used today. Three-dimensional cell culture is typically realized by the use of hydrogels. A hydrogel is a network of hydrophilic polymers, often used as a scaffold for 3-D cell culture. Due to its high water content, hydrogels provide environment similar to ECM of in vivo tissues. Several natural and synthetic hydrogels are commercially available. Natural hydrogels, such as collagen and Matrigel, have advantages that it closely mimics the chemical nature of in vivo tissues, while synthetic hydrogels such as poly ethylene glycol (PEG) are easy to use, and fine-tuning of its mechanical and chemical properties is possible.15 Three-dimensional organization is observed not only in regular tissues but also in a tumor mass. Vascularization inside a tumor mass is controlled by the delivery of proangiogenic factors and interaction with vascular cells, and has a significant effect on the tumor development and therapy.49 Combination of microfluidic culture models and 3-D hydrogel cell culture

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models is likely to provide a novel in vitro platform that mimics the tumor microenvironment and enable physiologically relevant testing of hypotheses and therapies.66

Mechanical Stimulus Cells are exposed to diverse mechanical stimuli in the natural tissue environment. In blood vessels, fluidic shear from the blood flow plays an important role in endothelial cell behavior, while compressive and tensile force also affects cell behavior. The fluidic shear stress is known to affect cell adhesion to surface, membrane permeability, and migration of cells.88 Microfluidics is an ideal tool for studying the effect of flow on cell biology, as it allows precise control and mathematical description of fluid flow.33 Several cell culture flow systems have been developed to study the effects of mechanical forces on endothelial cells. Conventional, macroscale devices employed parallel plate flow chamber design, but the major drawback is that it is difficult to test various conditions (cell type, surface property, flow rate, etc.) in a single experiment.86 On the other hand, microfluidics allows parallelization of multiple conditions more easily. For example, by manipulating the geometry and surface chemistry, cell adhesion in various shear forces and biochemistry were tested.41 The flow inside a microfluidic channel is determined by the volumetric flow rate and the geometry of the fluidic channel. Solving the Navier–Stokes equation gives the velocity profiles and pressure drops of the flow. In case of a flat parallel plate, the wall shear stress can be expressed by the following equation. s ¼ 6lQ=wh2 where Q is the flow rate, l is the viscosity, w is the width and h is the height of the channel. This equation is valid for a channel with a relatively flat geometry, that is, w is greater than h. In case of a channel with more square shapes (w = h), the wall effect becomes more significant, and the solution can be derived as follows.   2lQ m þ 1 s¼ ðn þ 1Þ wh2 m where m and n are empirical constants that depend on the aspect ratio a = h/w.60 It should be noted that the wall and the boundary layer effects become more pronounced as the system becomes smaller. Also these equations hold for a fully developed flow, and the entrance region should be considered separately. These equations describe the relationship between the shear stress and the channel geometry. Since microfabrication

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technique allows easy manipulation of channel geometry, it becomes possible to create multiple channels with desired fluidic properties. In addition to the fluidic shear, cells are also exposed to various non-shear forces coming from neighboring cells or ECM, such as tensile and compressive forces. For example, mechanical ventilation for a patient with respiratory disease causes alveolar epithelial cells in the lung to be exposed to a cyclic stretching and relaxation, resulting in mechanical stress and ventilator-induced lung injury.76 Takayama et al.13 developed a microfluidic alveolar model to study this mechanical stress on lung epithelial cells. Mechanical signals often originate from the interaction of cells and surrounding matrix. Diverse natural and synthetic hydrogels have been used to examine the mechanical properties of hydrogels and its effect on cells.15 Various experimental methods for determining the mechanical properties of hydrogels have been reviewed.30 Various microfluidic schemes have been developed to apply mechanical forces on cultured cells or hydrogel scaffolds and examine its effect. Optical laser tweezers have been integrated with a microfluidic device to apply tensile and compressive forces on a single cell. Beads functionalized with ECM proteins were manipulated with the optical tweezers to apply forces to ECM–integrin–cytoskeleton linkage.20 Such a system was successfully used to study focal adhesion recruitment. A valve-based microfluidic system was used to apply compression on neuron axons.21 Deformation of the axons was observed while applying force using regulated compressed gas. In another study, shear forces and non-shear, compressive forces were combined to mimic the physiological environment of the lacunar-canalicular porosity of bone.54

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responsible for the metabolism of drugs before they reach the rest of the body.7 The interaction between the liver and the target organ can have a significant effect on the pharmacokinetics and the effect of drugs. When studying the mechanism of diseases, reductionism has been the predominant approach. In other words, a biological system is divided into smaller systems and each system is examined separately. For example, an organism is broken down to organs, which are broken down to tissues, to cells, to proteins, and genes, etc.5 However, as Aristotle noted in the Metaphysica, ‘‘the whole is different from the sum of its parts’’. When applied to a living organism, the dynamic interaction between different organs certainly adds complexity to the whole system, and the whole-body response can often be explained only in terms of ‘‘orchestration’’ of multiple organs. In this perspective, systems approach might be better suited for understanding the dynamics of multi-organ interactions. Microfluidics has been used to capture the multiorgan interaction in the body. Termed as ‘‘cell culture analog’’, or ‘‘human-on-a-chip’’, this device allows investigation of whole-body response to drugs by culturing multiple cell types on a single device, and connecting those cells with fluidic channels mimicking the blood circulation.62,68–70,80 As will be described in more details later, they clearly demonstrate the possibility of using microfluidics to mimic the whole-body response. Another important aspect of such multiorgan interaction is that it is a dynamic process, changing with time. Such time-dependent processes can be reproduced with microfluidic systems. A good example is the investigation of the dynamics of genetic network in a single cell.4 Co-Culture

Dynamics of Multi-Organ Interaction A fundamental difference between in vitro cell culture and the whole body is the presence of multi-organ interactions. Homeostasis is orchestrated by the complex interaction between different organs in the body. A prominent example is endocrine signaling, where organs secret hormones into the bloodstream to regulate the body. In response to an increase in the elevation in glycemia, insulin is secreted from the pancreas, and transported via the circulatory system to other organs including the liver, fat, and the muscle. Insulin then binds to cell receptors and stimulates glucose disposal.52 Another example is the first-pass metabolism of drugs, which refers to the metabolism of orally administered drugs before entering the systemic circulation. This process is mostly mediated by the gut and the liver. The liver is strategically placed after the digestive tract through the portal vein, and is

In natural tissue environment, different cell types co-exist while communicating with one another. To mimic this, cells are often co-cultured with different cell types. For example, hepatocytes have been cultured with hepatic Kupffer cells to examine the effect of intercellular communication on xenobiotic metabolism in the liver.48 However, the only controllable parameter in the conventional co-culture model is the ratio between different cell types. This might not fully capture the aspect of the natural tissue environment, which has complex hierarchical structures. Bhatia and co-workers31 reported using stencil-based method to micropattern hepatocytes and stromal cells to create functional subunits of the liver. The enhancement in the liver specific function was observed when hepatocytes were co-cultured with fibroblasts. More importantly, the micropatterned co-culture outperformed randomly distributed co-culture model. By changing

Microtechnology for Mimicking In Vivo Tissue Environment

the relative size of each cell populations, they were able to find the optimal combination where the liver-specific function was maximized. Co-culture models have also been used in a microfluidic device. To develop an in vitro absorption model of iron, mucus producing cells (HT29-MTX) were co-cultured with Caco-2 cell line, which is frequently used for drug absorption studies.45 This co-culture model was later used in a microfluidic setting to examine the first-pass metabolism of orally delivered drugs, acetominophen.44 Although less common than co-culture models, triple cell culture models have been examined. Lehmann et al.38 reported triple cell culture model of epithelial cells, macrophages, and dendritic cells to study the interaction of xenobiotics with those cells. METHODS IN MICROTECHNOLOGY Photolithography and Soft Lithography Photolithography comprises the basis of most of microtechnology used today. It uses photoresist, a

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photo-sensitive material that changes its property upon exposure to UV light, and a photomask to create arbitrary patterns on the photoresist coated on the substrate (Fig. 2a). Subsequent etching, development, or deposition results in the fabrication of desired structures on the substrate.55 Although conventional silicon etching method has been widely used in microfluidics, it is relatively expensive and fabrication process is complex and requires a clean room. Soft lithography uses polydimethylsiloxane (PDMS), an elastomeric material which is inexpensive and easy to use, transparent for optical examination, gas permeable which is important for cell culture, known to be biocompatible and nontoxic to cells. However, recent studies show that PDMS carries a few potential problems. The most well-known problem is non-specific binding of molecules to its surface. Due to its high hydrophobicity, hydrophobic small molecules are affected most by this problem.73 Another is the potential leaching out of uncured PDMS monomers, and its effect on cells.87 Although PDMS is thought to be non-toxic to cells at

FIGURE 2. Microscale technologies (a) photolithography, (b) pneumatic flow control, (c) stencil-based cell patterning, and (d) hydrogel microfabrication.

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least in the short term, its long-term effect on cells, especially non-lethal, subtle changes, still has to be evaluated carefully. Fluidic Manipulation Microfluidics manipulates tiny quantity of liquid (from nanoliters to microliters) in channels with dimensions of tens of micrometers.83 While microfluidics offers many advantages such as small sample consumption, high sensitivity, resolution, and low cost, having to deal with a small quantity of liquid requires novel equipment to enable precise control and manipulation of the liquid. In this perspective, microfluidic valves and pumps have been of great interest to researchers. PDMS has been a favorite material, as its flexible nature allows fabrication of built-in pumps and valves through multi-layer lithography (Fig. 2b).90 Pioneered by Quake lab, the basic principle is to insert or withdraw air or liquid into PDMS channels (control layer) to either block or open the liquid channels. Combination with computer-controlled solenoid enabled fabrication of a micropump with a relatively wide range of pumping rates.89 Mixing is another area of interest in microfluidics. The small size in microfluidics naturally results in a laminar flow with diffusion-dominating transport, which has been used extensively for precise control of the flow. While mixing is intentionally minimized in this setting, it might not be appropriate in the cases where mixing is desired. Controlled mixing is especially important in microreactor applications, such as chemical synthesis, DNA analysis, and enzyme assays.26 Microscale magnetic particle has been used in a microfluidic system to enhance mixing.37 Natural convection was induced in a microfluidic device by incorporating two alternating heating element into a membrane.34

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selectively block adhesion of specific cells. Removal of the stencil and subsequent adhesion allows patterning of different cell types in desired locations. Stamping or microcontact printing uses a PDMS stamp to create patterns of molecules that promotes or inhibits adhesion of cells. This technique is also relatively cheap and easy to use once the stamp has been made. All these methods are typically applied to 2-D surfaces, and recent efforts have been directed to patterning cells in 3-D space.18 Hydrogel Microfabrication Hydrogels are used to provide cells with 3-D support that mimics natural ECM environment. Earlier studies were limited to using simple, macroscale shapes of hydrogel scaffold, such as cylindrical or cubic shapes with relatively large sizes. Recent efforts have been directed to fabricating hydrogels to mimic microscale structures of target organ tissues. Several methods have been developed, such as micromolding, photolithography using UV-curable polymers, direct printing, and bottom-up approach.29,75 These methods allow fabrication of microscale shapes in hydrogels. Cellladen hydrogels can also be fabricated by mixing cells with hydrogels before the gelation process. Fabrication of complex 3-D shapes has also been reported. Sung et al.72 have developed a novel protocol for fabricating high aspect-ratio structures with a curvature using collagen, and demonstrated fabrication of hydrogel structure mimicking the intestinal villi. The curved, finger-like shape of the intestinal villi makes it difficult to mimic using conventional hydrogel fabrication methods. Another notable example is a bottom-up, modular approach to create 3-D structure. This is potentially useful for mimicking the structure of tissue that has repeating functional units. For example, rodshaped collagen gels that were seeded with hepatocytes on the inside and endothelial cells on the surface were packed within a bioreactor.46

Cell Patterning As mentioned earlier, the natural tissue consists of several cell types, organized in a hierarchical manner. The ability to located cells in desired locations enables mimicking the complex cellular organization of the natural tissues. Several cell patterning techniques have been developed, such as photolithographic patterning,6 stencil-based patterning,85 stamping,51 laser-guided cell patterning,59 electrochemical method,17 and microfluidics.74 Photographic patterning creates micropatterns using light and a photomask to selectively coat the surface with cell-friendly molecules such as fibronectin and collagen. While photolithographic patterning requires expensive cleanroom facilities, stencil-based patterning is cheaper and easier. A PDMS stencil has been used to

APPLICATIONS Microscale Biomimetic Tissues Application of microtechnology in biomedical field is diverse. Here we will focus on the effort to develop in vitro systems mimicking specific organs. Diverse approaches have been taken to fabricate microscale systems mimicking the liver, kidney, lung, blood vessels, etc. The liver is responsible for biotransformation and detoxification of xenobiotics. Due to its importance, several in vitro systems have been developed to reproduce the liver metabolism, which are summarized in

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several review articles.3,53,56 These attempts go beyond simply culturing hepatocytes in vitro, and try to reproduce the microenvironment of the liver tissue. Earlier attempts tried to mimic the liver tissue by culturing hepatocytes in collagen matrix.11 More recent attempts utilized microtechnology to reproduce the 3-D cell culture with fluid flow,63 zonation of oxygen concentration,1 co-culture environment,31 or perfuse liver tissue slices.78 In these studies, enhanced hepatic functions were observed with hepatocytes cultured in the devices. The kidney plays a major role in eliminating waste metabolites and exogenous compounds from the body. Reproducing the kidney function poses an interesting challenge as it involves fluid flow and transport of small molecules. A simple hemodialysis device has been used for patients with renal failure,23 but it can result in sudden change in blood chemistry and volume due to non-physiological flow rate. Microfabrication technique was used to create a controlled laminar flow to enable membrane-less dialysis.39 Leclerc and co-workers57 has developed a microscale renal chip to simulate the mass transport in the kidney by reproducing the glomerular apical and basolateral sides (Fig. 2). The lung is responsible for the exchange of oxygen and carbon dioxide between the atmosphere and bloodstream. The exchange of gas occurs in the subunits of the lung called the alveoli. Hollow fiber membranes have been used to simulate the gas exchange in the lung.84 A more recent, microtechnology-based approach includes using soft-lithography to create 3-D array of blood microchannels and gas pathways to simulate the natural lung.8 Huh et al.22, reconstituted the alveolar-capillary interface and simulated the mechanical movement of the lung tissue by applying vacuum to PDMS channels. Exposure of the artificial lung-on-a-chip to nanoparticles revealed that the cyclic mechanical strain accentuates toxic and inflammatory responses of the lung. The process of oral absorption is an important factor that determines the pharmacokinetics of drugs. The most widely used in vitro model of oral absorption is the Caco-2 cell model.2 After differentiation, Caco-2 cells form polar, enterocyte-like epithelial cell layers with tight junctions and microvilli. It is thought that drugs can be absorbed via several pathways including passive transport which requires a concentration gradient, and active, carrier-mediated transport. Differentiated Caco-2 cells are thought to provide at least some of these transport pathways.2 To reproduce the absorption process in vitro, two separate compartments (apical and basolateral sides) separated by semipermeable membrane need to be created. In conventional drug absorption study, a confluent layer of

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Caco-2 cells is cultured on a semipermeable membrane, and the transport of drugs through the membrane is measured. However, 2-D monolayer culture of Caco-2 cells is not an accurate representation of intestinal epithelium. The intestinal epithelium contains finger-like projections called villi, which increases the absorptive surface area. It is thought that this 3-D geometry affects the absorption kinetics.2 To reproduce the 3-D geometry of intestinal villi, a novel fabrication method was developed.72 Collagen was fabricated to mimic the size and density of human intestinal villi, and Caco-2 cells were cultured on the hydrogel scaffold. Cells grew conforming to the hydrogel structure, forming 3-D shapes similar to the natural intestinal villi. Another limitation of current Caco-2 model is that it is a static system, whereas the natural intestinal tissue is more dynamic with the blood flow and the peristalsis. A two-layer microfluidic device can simulate the process better since it can reproduce the fluid flow in the apical and basolateral sides. The vascular network is simulated by culturing endothelial cells on a porous membrane.65 Such systems are used to assess the change in the permeability of the endothelial lining in response to drugs. A challenge in the case of reproducing the vascular network is how to fabricate the round structure of the blood vessels. Hollow collagen endothelial vessels were formed by crosslinking around needles that were subsequently removed. Novel microfabrication methods have been developed to create rectangular, trapezoidal, and circular cross-sectional profiles.9,14,16 Mimicking Whole-Body Response and Dynamics A true complexity of the human body comes from the dynamic interactions between organs. For example, an orally administered drug can go through absorption in the GI tract, metabolism in the liver, distribution throughout the body and elimination through the kidney or the liver. These processes, collectively known as absorption, distribution, metabolism and elimination (ADME) contribute to the pharmacokinetic profile of drugs. A mathematical approach is available to simulate such processes, called physiologically based pharmacokinetic (PBPK) modeling, but this approach has several limitations. The most serious one is the difficulty associated with obtaining a reliable set of data to build an accurate model.68 Using microfluidics, a physical replica of a pharmacokinetic model has been developed.62 This microchip contains separate cell culture microchambers representing different organs, connected with fluidic channels to simulate the blood flow. Termed cell culture analogs or human-on-a-chip, such in vitro platforms allow direct assessment of hypotheses in the mathematical models

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(Fig. 2). It has been shown that this device can be used to reproduce the metabolism-dependent toxicity of naphthalene and anti-cancer drugs.70,80 More recently, the device has been modified to allow 3-D cell culture inside the device for more realistic tissue environment.69 Absorption and subsequent metabolism of acetaminophen has been simulated by combining an in vitro model of gastrointestinal (GI) tract with a microscale cell culture analog representing the body.44 Metabolism by the intestinal epithelium cell line and liver cells was observed. Similar attempts to reproduce multi-organ interaction or metabolism-dependent toxicity of drugs have been made. A sol–gel human liver microsome (HLM) bioreactor was coupled with a cell culture chamber array.43 Drugs are metabolized by HLM, flow into the cell culture chamber array, where the cytotoxicity of the metabolites and the drugs is assessed. A microarray of sol–gel encapsulated P450 enzymes has been developed.36 Target drugs are metabolized by the Metachip (a microarray of P450 enzymes encapsulated by hydrogels), which is superimposed on a Datachip (a microarray of cell cultures) to assess the toxicity of the metabolites. Ahluwalia and co-workers,81 has developed a multicompartmental modular bioreactor to allow multiple cell types to be cultured on a single device. Each compartment is connected with channels or tubes to simulate the blood flow. Using this device, a cross-talk between hepatocytes and adipose tissue was reproduced.19 A sequential perfusion of the intestinal and liver slices has been realized using twocompartment perfusion system to mimic the first-pass metabolism (Fig. 2).78 When the slices were exposed to the primary bile acid, expression of fibroblast growth factor 15 in the intestinal slice was induced, which subsequently down-regulated cytochrome P450 enzyme in the liver slice. This observation successfully demonstrated the interplay between the two organs.

REMAINING CHALLENGES AND CONCLUSION The greatest challenge of current microtechnology is how to enable user-friendly, high-throughput implementation. The major reason why we are not seeing the direct application of these technologies in biomedical field is because it requires highly skilled personnel to operate the devices. A pressure-driven flow by using a peristaltic or syringe pump is the easiest way to provide a consistent flow, but requires fluidic connections via tubes, which can make high-throughput implementation challenging. Other forms of flow, such as electroosmotic flow, are available but it also requires certain equipment coupled with the microfluidic devices. In this perspective, recent development in pump-less

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microfluidic systems and high-throughput implementation is encouraging.47 Pump-less microfluidic systems uses passive pumping method, originating from the difference in the surface tension of liquid drops at the inlet and outlet ports of the device. Another useful method is the use of gravity-induced flow. By tilting the device, a height difference between the outlet and the inlet causes a gravity-driven flow. The flow rate depends on the height difference and the geometry of the channel.50 Recently, gravity-induced flow was utilized to develop a microfluidic device for testing the effect of anti-cancer agents.69 Such passive pumping methods eliminate the need of a pump or expensive equipment to be connected with microfluidic devices, facilitating high-throughput implementation (Fig. 3). Microfluidic devices typically consist of fluidic channels with widths of several micrometers to hundred micrometers. Such narrow channels are prone to the creation of air bubbles inside the channels. Bubbles trapped inside the channels can be problematic because they can block the channels, distorting the desired flow patterns, or damaging cells by shearing at the air– liquid interface. Bubble traps have been invented to prevent such problems.71 Most bubble traps utilize the buoyancy of air bubbles, but a few other methods have been devised, such as pressure-driven bubble removal, or 2-layer structure with pneumatic chamber to pull the bubble through a PDMS membrane.27,64 Ideal bubble traps should allow easy integration with microfluidic devices, efficiently remove air bubbles, and should not distort or affect the main fluidic circuit. An important aspect that has often been overlooked is the fact that cells cultured in microdevices often behave differently from when they are cultured in conventional, macroscale systems. The growth kinetics and protein expression profiles in a microfluidic system and a conventional cell culture system have been compared.87 Differences in proliferation, glucose metabolism, signaling pathway activation and protein expression have been observed. The reasons for such a difference are thought to be the edge effect, fluidic shear, and surface area-to-volume ratio, but detailed mechanisms still need to be elucidated. Young and Beebe87 has provided the concept of effective culture volume (ECV), effective culture time (ECT), and a critical perfusion rate (CPR) to quantify the differences caused by the small scale of microdevices. The advantages of microtechnology are clear. We are now seeing many examples of microsystems mimicking the microenvironment of natural tissues. These systems allow studying biological systems in a way that has not been possible with conventional, macroscale systems. Although this technology still has several limitations, recent progress in the field has been remarkable and we believe that in near future we will

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FIGURE 3. Applications of microscale technology (a) renal microchip, figure was drawn based on ref 57, (b) microfluidic device for intestinal and liver tissue slice, reproduced with permission from Royal Society of Chemistry,78, and (c) microscale cell culture analog (human-on a-chip).

see increased direct application of this technology in biology and medicine.

ACKNOWLEDGMENTS This work was supported by Army Corp of Engineers (CERL, W9132T-07), Nanobiotechnology center (NBTC), National Research Foundation of Korea (NRF, Grant no. 2011-0013862), Hongik University new faculty research support fund, and 2011 Hongik University Research Fund.

REFERENCES 1

Allen, J. W., S. R. Khetani, and S. N. Bhatia. In vitro zonation and toxicity in a hepatocyte bioreactor. Toxicol. Sci. 84(1):110–119, 2005. 2 Artursson, P., K. Palm, and K. Luthman. Caco-2 monolayers in experimental and theoretical predictions of drug transport. Adv. Drug Deliv. Rev. 46(1–3):27–43, 2001. 3 Baudoin, R., A. Corlu, L. Griscom, C. Legallais, and E. Leclerc. Trends in the development of microfluidic cell biochips for in vitro hepatotoxicity. Toxicol. In Vitro 21(4):535–544, 2007.

4

Bennett, M. R., and J. Hasty. Microfluidic devices for measuring gene network dynamics in single cells. Nat. Rev. Genet. 10(9):628–638, 2009. 5 Bergman, R. N. Orchestration of glucose homeostasis: from a small acorn to the California oak. Diabetes 56(6):1489–1501, 2007. 6 Bhatia, S. N., M. L. Yarmush, and M. Toner. Controlling cell interactions by micropatterning in co-cultures: hepatocytes and 3T3 fibroblasts. J. Biomed. Mater. Res. 34(2):189–199, 1997. 7 Brandon, E. F., C. D. Raap, I. Meijerman, J. H. Beijnen, and J. H. Schellens. An update on in vitro test methods in human hepatic drug biotransformation research: pros and cons. Toxicol. Appl. Pharmacol. 189(3):233–246, 2003. 8 Burgess, K. A., H. H. Hu, W. R. Wagner, and W. J. Federspiel. Towards microfabricated biohybrid artificial lung modules for chronic respiratory support. Biomed. Microdevices 11(1):117–127, 2009. 9 Camp, J. P., T. Stokol, and M. L. Shuler. Fabrication of a multiple-diameter branched network of microvascular channels with semi-circular cross-sections using xenon difluoride etching. Biomed. Microdevices 10(2):179–186, 2008. 10 Cukierman, E., R. Pankov, D. R. Stevens, and K. M. Yamada. Taking cell–matrix adhesions to the third dimension. Science 294(5547):1708–1712, 2001. 11 De Smet, K., T. Bruning, M. Blaszkewicz, H. M. Bolt, A. Vercruysse, and V. Rogiers. Biotransformation of trichloroethylene in collagen gel sandwich cultures of rat hepatocytes. Arch. Toxicol. 74(10):587–592, 2000.

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J. H. SUNG

AND

Dittrich, P. S., and A. Manz. Lab-on-a-chip: microfluidics in drug discovery. Nat. Rev. Drug Discov. 5(3):210–218, 2006. 13 Douville, N. J., P. Zamankhan, Y. C. Tung, R. Li, B. L. Vaughan, C. F. Tai, J. White, P. J. Christensen, J. B. Grotberg, and S. Takayama. Combination of fluid and solid mechanical stresses contribute to cell death and detachment in a microfluidic alveolar model. Lab Chip 11(4):609–619, 2011. 14 Fidkowski, C., M. R. Kaazempur-Mofrad, J. Borenstein, J. P. Vacanti, R. Langer, and Y. Wang. Endothelialized microvasculature based on a biodegradable elastomer. Tissue Eng. 11(1–2):302–309, 2005. 15 Geckil, H., F. Xu, X. Zhang, S. Moon, and U. Demirci. Engineering hydrogels as extracellular matrix mimics. Nanomedicine (Lond.) 5(3):469–484, 2010. 16 Golden, A. P., and J. Tien. Fabrication of microfluidic hydrogels using molded gelatin as a sacrificial element. Lab Chip 7(6):720–725, 2007. 17 Guillaume-Gentil, O., M. Gabi, M. Zenobi-Wong, and J. Voros. Electrochemically switchable platform for the micro-patterning and release of heterotypic cell sheets. Biomed. Microdevices 13(1):221–230, 2011. 18 Guillotin, B., and F. Guillemot. Cell patterning technologies for organotypic tissue fabrication. Trends Biotechnol. 29(4):183–190, 2011. 19 Guzzardi, M. A., C. Domenici, and A. Ahluwalia. Metabolic control through hepatocyte and adipose tissue crosstalk in a multicompartmental modular bioreactor. Tissue Eng. A 17(11–12):1635–1642, 2011. 20 Honarmandi, P., H. Lee, M. J. Lang, and R. D. Kamm. A microfluidic system with optical laser tweezers to study mechanotransduction and focal adhesion recruitment. Lab Chip 11(4):684–694, 2011. 21 Hosmane, S., A. Fournier, R. Wright, L. Rajbhandari, R. Siddique, I. H. Yang, K. T. Ramesh, A. Venkatesan, and N. Thakor. Valve-based microfluidic compression platform: single axon injury and regrowth. Lab Chip 11(22):3888–3895, 2011. 22 Huh, D., B. D. Matthews, A. Mammoto, M. MontoyaZavala, H. Y. Hsin, and D. E. Ingber. Reconstituting organ-level lung functions on a chip. Science 328(5986): 1662–1668, 2010. 23 Humes, H. D., W. H. Fissell, and K. Tiranathanagul. The future of hemodialysis membranes. Kidney Int. 69(7):1115– 1119, 2006. 24 Ismagilov, R. F., and M. M. Maharbiz. Can we build synthetic, multicellular systems by controlling developmental signaling in space and time? Curr. Opin. Chem. Biol. 11(6):604–611, 2007. 25 Janmey, P. A., and C. A. McCulloch. Cell mechanics: integrating cell responses to mechanical stimuli. Annu. Rev. Biomed. Eng. 9:1–34, 2007. 26 Jeong, G. S., S. Chung, C. B. Kim, and S. H. Lee. Applications of micromixing technology. Analyst 135(3):460–473, 2010. 27 Kang, J. H., Y. C. Kim, and J. K. Park. Analysis of pressure-driven air bubble elimination in a microfluidic device. Lab Chip 8(1):176–178, 2008. 28 Keenan, T. M., and A. Folch. Biomolecular gradients in cell culture systems. Lab Chip 8(1):34–57, 2008. 29 Khademhosseini, A., and R. Langer. Microengineered hydrogels for tissue engineering. Biomaterials 28(34):5087– 5092, 2007. 30 Khaleque, T., S. Abu-Salih, J. R. Saunders, and W. Moussa. Experimental methods of actuation, characterization and

M. L. SHULER prototyping of hydrogels for bioMEMS/NEMS applications. J. Nanosci. Nanotechnol. 11(3):2470–2479, 2011. 31 Khetani, S. R., and S. N. Bhatia. Microscale culture of human liver cells for drug development. Nat. Biotechnol. 26(1):120–126, 2008. 32 Kim, S., H. J. Kim, and N. L. Jeon. Biological applications of microfluidic gradient devices. Integr. Biol. (Camb.) 2(1112):584–603, 2010. 33 Kim, L., M. D. Vahey, H. Y. Lee, and J. Voldman. Microfluidic arrays for logarithmically perfused embryonic stem cell culture. Lab Chip 6(3):394–406, 2006. 34 Kim, S. J., F. Wang, M. A. Burns, and K. Kurabayashi. Temperature-programmed natural convection for micromixing and biochemical reaction in a single microfluidic chamber. Anal. Chem. 81(11):4510–4516, 2009. 35 Lee, K., C. Kim, B. Ahn, R. Panchapakesan, A. R. Full, L. Nordee, J. Y. Kang, and K. W. Oh. Generalized serial dilution module for monotonic and arbitrary microfluidic gradient generators. Lab Chip 9(5):709–717, 2009. 36 Lee, M. Y., R. A. Kumar, S. M. Sukumaran, M. G. Hogg, D. S. Clark, and J. S. Dordick. Three-dimensional cellular microarray for high-throughput toxicology assays. Proc. Natl. Acad. Sci. USA 105(1):59–63, 2008. 37 Lee, S. H., D. van Noort, J. Y. Lee, B. T. Zhang, and T. H. Park. Effective mixing in a microfluidic chip using magnetic particles. Lab Chip. 9(3):479–482, 2009. 38 Lehmann, A. D., N. Daum, M. Bur, C. M. Lehr, P. Gehr, and B. M. Rothen-Rutishauser. An in vitro triple cell co-culture model with primary cells mimicking the human alveolar epithelial barrier. Eur. J. Pharm. Biopharm. 77(3):398–406, 2011. 39 Leonard, E. F., S. Cortell, and N. G. Vitale. Membraneless dialysis—is it possible? Contrib. Nephrol. 149:343–353, 2005. 40 Li Jeon, N., H. Baskaran, S. K. Dertinger, G. M. Whitesides, L. Van de Water, and M. Toner. Neutrophil chemotaxis in linear and complex gradients of interleukin-8 formed in a microfabricated device. Nat. Biotechnol. 20(8):826–830, 2002. 41 Lu, H., L. Y. Koo, W. M. Wang, D. A. Lauffenburger, L. G. Griffith, and K. F. Jensen. Microfluidic shear devices for quantitative analysis of cell adhesion. Anal. Chem. 76(18):5257–5264, 2004. 42 Lucchetta, E. M., J. H. Lee, L. A. Fu, N. H. Patel, and R. F. Ismagilov. Dynamics of Drosophila embryonic patterning network perturbed in space and time using microfluidics. Nature 434(7037):1134–1138, 2005. 43 Ma, B., G. Zhang, J. Qin, and B. Lin. Characterization of drug metabolites and cytotoxicity assay simultaneously using an integrated microfluidic device. Lab Chip 9(2):232– 238, 2009. 44 Mahler, G. J., M. B. Esch, R. P. Glahn, and M. L. Shuler. Characterization of a gastrointestinal tract microscale cell culture analog used to predict drug toxicity. Biotechnol. Bioeng. 104(1):193–205, 2009. 45 Mahler, G. J., M. L. Shuler, and R. P. Glahn. Characterization of Caco-2 and HT29-MTX cocultures in an in vitro digestion/cell culture model used to predict iron bioavailability. J. Nutr. Biochem. 20(7):494–502, 2009. 46 McGuigan, A. P., and M. V. Sefton. Vascularized organoid engineered by modular assembly enables blood perfusion. Proc. Natl. Acad. Sci. USA 103(31):11461–11466, 2006. 47 Meyvantsson, I., J. W. Warrick, S. Hayes, A. Skoien, and D. J. Beebe. Automated cell culture in high density tubeless microfluidic device arrays. Lab Chip 8(5):717–724, 2008.

Microtechnology for Mimicking In Vivo Tissue Environment 48

Milosevic, N., H. Schawalder, and P. Maier. Kupffer cellmediated differential down-regulation of cytochrome P450 metabolism in rat hepatocytes. Eur. J. Pharmacol. 368(1): 75–87, 1999. 49 Moon, J. J., and J. L. West. Vascularization of engineered tissues: approaches to promote angio-genesis in biomaterials. Curr. Top Med. Chem. 8(4):300–310, 2008. 50 Morier, P., C. Vollet, P. E. Michel, F. Reymond, and J. S. Rossier. Gravity-induced convective flow in microfluidic systems: electrochemical characterization and application to enzyme-linked immunosorbent assay tests. Electrophoresis 25(21–22):3761–3768, 2004. 51 Mrksich, M., L. E. Dike, J. Tien, D. E. Ingber, and G. M. Whitesides. Using microcontact printing to pattern the attachment of mammalian cells to self-assembled monolayers of alkanethiolates on transparent films of gold and silver. Exp. Cell Res. 235(2):305–313, 1997. 52 Musi, N., and L. J. Goodyear. Insulin resistance and improvements in signal transduction. Endocrine 29(1):73– 80, 2006. 53 Nahmias, Y., F. Berthiaume, and M. L. Yarmush. Integration of technologies for hepatic tissue engineering. Adv. Biochem. Eng. Biotechnol. 103:309–329, 2007. 54 Orr, D. E., and K. J. Burg. Design of a modular bioreactor to incorporate both perfusion flow and hydrostatic compression for tissue engineering applications. Ann. Biomed. Eng. 36(7):1228–1241, 2008. 55 Park, T. H., and M. L. Shuler. Integration of cell culture and microfabrication technology. Biotechnol. Prog. 19(2): 243–253, 2003. 56 Pelkonen, O., and M. Turpeinen. In vitro–in vivo extrapolation of hepatic clearance: biological tools, scaling factors, model assumptions and correct concentrations. Xenobiotica 37(10–11):1066–1089, 2007. 57 Ramello, C., P. Paullier, A. Ould-Dris, M. Monge, C. Legallais, and E. Leclerc. Investigation into modification of mass transfer kinetics by acrolein in a renal biochip. Toxicol. In Vitro 25(5):1123–1131, 2011. 58 Saadi, W., S. W. Rhee, F. Lin, B. Vahidi, B. G. Chung, and N. L. Jeon. Generation of stable concentration gradients in 2D and 3D environments using a microfluidic ladder chamber. Biomed. Microdevices 9(5):627–635, 2007. 59 Schiele, N. R., D. T. Corr, Y. Huang, N. A. Raof, Y. Xie, and D. B. Chrisey. Laser-based direct-write techniques for cell printing. Biofabrication 2(3):032001, 2010. 60 Shah, R. K., and A. L. London, Laminar Flow Forced Convection in Ducts: A Source Book for Compact Heat Exchanger Analytical Data. Advances in Heat Transfer Supplement. New York: Academic Press, xiv, 477 pp., 1978. 61 Sharma, R. I., and J. G. Snedeker. Biochemical and biomechanical gradients for directed bone marrow stromal cell differentiation toward tendon and bone. Biomaterials 31(30):7695–7704, 2010. 62 Sin, A., K. C. Chin, M. F. Jamil, Y. Kostov, G. Rao, and M. L. Shuler. The design and fabrication of three-chamber microscale cell culture analog devices with integrated dissolved oxygen sensors. Biotechnol. Prog. 20(1):338–345, 2004. 63 Sivaraman, A., J. K. Leach, S. Townsend, T. Iida, B. J. Hogan, D. B. Stolz, R. Fry, L. D. Samson, S. R. Tannenbaum, and L. G. Griffith. A microscale in vitro physiological model of the liver: predictive screens for drug metabolism and enzyme induction. Curr. Drug Metab. 6(6):569–591, 2005.

64

1299

Skelley, A. M., and J. Voldman. An active bubble trap and debubbler for microfluidic systems. Lab Chip 8(10):1733– 1737, 2008. 65 Stoltz, J. F., S. Muller, A. Kadi, V. Decot, P. Menu, and D. Bensoussan. Introduction to endothelial cell biology. Clin. Hemorheol. Microcirc. 37(1–2):5–8, 2007. 66 Stroock, A. D., and C. Fischbach. Microfluidic culture models of tumor angiogenesis. Tissue Eng. A 16(7):2143– 2146, 2010. 67 Sundararaghavan, H. G., G. A. Monteiro, B. L. Firestein, and D. I. Shreiber. Neurite growth in 3D collagen gels with gradients of mechanical properties. Biotechnol. Bioeng. 102(2):632–643, 2009. 68 Sung, J. H., M. B. Esch, and M. L. Shuler. Integration of in silico and in vitro platforms for pharmacokinetic-pharmacodynamic modeling. Expert Opin. Drug Metab. Toxicol. 6(9):1063–1081, 2010. 69 Sung, J. H., C. Kam, and M. L. Shuler. A microfluidic device for a pharmacokinetic-pharmacodynamic (PK-PD) model on a chip. Lab Chip 10(4):446–455, 2010. 70 Sung, J. H., and M. L. Shuler. A micro cell culture analog (microCCA) with 3-D hydrogel culture of multiple cell lines to assess metabolism-dependent cytotoxicity of anti-cancer drugs. Lab Chip 9(10):1385–1394, 2009. 71 Sung, J. H., and M. L. Shuler. Prevention of air bubble formation in a microfluidic perfusion cell culture system using a microscale bubble trap. Biomed. Microdevices 11(4):731–738, 2009. 72 Sung, J. H., J. Yu, D. Luo, M. L. Shuler, and J. C. March. Microscale 3-D hydrogel scaffold for biomimetic gastrointestinal (GI) tract model. Lab Chip 11(3):389–392, 2011. 73 Toepke, M. W., and D. J. Beebe. PDMS absorption of small molecules and consequences in microfluidic applications. Lab Chip 6(12):1484–1486, 2006. 74 Torisawa, Y. S., B. Mosadegh, G. D. Luker, M. Morell, K. S. O’Shea, and S. Takayama. Microfluidic hydrodynamic cellular patterning for systematic formation of coculture spheroids. Integr. Biol. (Camb.) 1(11–12):649–654, 2009. 75 Tsang, V. L., and S. N. Bhatia. Fabrication of threedimensional tissues. Adv. Biochem. Eng. Biotechnol. 103: 189–205, 2007. 76 Tschumperlin, D. J., and S. S. Margulies. Equibiaxial deformation-induced injury of alveolar epithelial cells in vitro. Am. J. Physiol. 275(6 Pt 1):L1173–L1183, 1998. 77 Tse, J. R., and A. J. Engler. Stiffness gradients mimicking in vivo tissue variation regulate mesenchymal stem cell fate. PLoS One 6(1):e15978, 2011. 78 van Midwoud, P. M., M. T. Merema, E. Verpoorte, and G. M. Groothuis. A microfluidic approach for in vitro assessment of interorgan interactions in drug metabolism using intestinal and liver slices. Lab Chip 10(20):2778–2786, 2010. 79 Vickerman, V., J. Blundo, S. Chung, and R. Kamm. Design, fabrication and implementation of a novel multiparameter control microfluidic platform for three-dimensional cell culture and real-time imaging. Lab Chip 8(9):1468–1477, 2008. 80 Viravaidya, K., A. Sin, and M. L. Shuler. Development of a microscale cell culture analog to probe naphthalene toxicity. Biotechnol. Prog. 20(1):316–323, 2004. 81 Vozzi, F., J. M. Heinrich, A. Bader, and A. D. Ahluwalia. Connected culture of murine hepatocytes and HUVEC in a multicompartmental bioreactor. Tissue Eng. A 15(6):1291– 1299, 2009.

1300 82

J. H. SUNG

AND

Wang, F., V. M. Weaver, O. W. Petersen, C. A. Larabell, S. Dedhar, P. Briand, R. Lupu, and M. J. Bissell. Reciprocal interactions between beta1-integrin and epidermal growth factor receptor in three-dimensional basement membrane breast cultures: a different perspective in epithelial biology. Proc. Natl. Acad. Sci. USA 95(25):14821–14826, 1998. 83 Whitesides, G. M. The origins and the future of microfluidics. Nature 442(7101):368–373, 2006. 84 Wnek, G. E., and G. L. Bowlin. Encyclopedia of Biomaterials and Biomedical Engineering. New York: Marcel Dekker, 2004. 85 Wright, D., B. Rajalingam, S. Selvarasah, M. R. Dokmeci, and A. Khademhosseini. Generation of static and dynamic patterned co-cultures using microfabricated parylene-C stencils. Lab Chip 7(10):1272–1279, 2007.

M. L. SHULER 86

Xiao, Y., and G. A. Truskey. Effect of receptor-ligand affinity on the strength of endothelial cell adhesion. Biophys. J. 71(5):2869–2884, 1996. 87 Young, E. W., and D. J. Beebe. Fundamentals of microfluidic cell culture in controlled microenvironments. Chem. Soc. Rev. 39(3):1036–1048, 2010. 88 Young, E. W., and C. A. Simmons. Macro- and microscale fluid flow systems for endothelial cell biology. Lab Chip 10(2):143–160, 2010. 89 Zhang, W., S. Lin, C. Wang, J. Hu, C. Li, Z. Zhuang, Y. Zhou, R. A. Mathies, and C. J. Yang. PMMA/PDMS valves and pumps for disposable microfluidics. Lab Chip 9(21):3088–3094, 2009. 90 Zheng, Y., W. Dai, and H. Wu. A screw-actuated pneumatic valve for portable, disposable microfluidics. Lab Chip 9(3):469–472, 2009.