Although enabling, conventional analytical technologies do not allow for an integrated handling, environmental control and analysis performed autonomously and with minimal operator supervision. These limitations can be, however, prospectively superseded by implementing technological advances recently made in the field of miniaturized LOC technologies that exploit laminar flow at the microscale (27, 28). Microfluidics is aimed at manipulating liquids at ultralow volumes in circuitries of small channels that have a cross-sectional area below a square millimeter (29, 30). As described by the dimensionless Reynolds number (Re), fluid flow in microfluidic channels is laminar and largely dominated by viscous forces (31, 32). Under these conditions, the fluid has no inertia. This enables accurate delivery of drugs, both spatially and temporally (33). The dawn of microfluidic chip-based technologies and its integration into the design of micro-total analysis systems (μTAS) heralds one of the most adventurous avenues to address the inherent complexity of cellular systems, with massive experimental parallelization and bioanalysis at the single cell level (28, 34). Importantly, the transfer of traditional bioanalytical methods to a microfabricated format provides a means to increase the resolution of bioanalysis, increase sampling throughput, and also allows the design of automated systems with innovative functionalities (28, 32, 35).
Not surprisingly, implementation of miniaturized systems for in situ analysis of small model organisms is attracting a rapidly growing interest. Recently, mushrooming reports have shown automated manipulation and immobilization predominantly of micron-sized organisms like C. elegans and D. melanogaster (36–38). Development of mesofluidic chip-based devices able to manipulate sub-millimeter scale organisms such as zebrafish and Xenopus eggs, embryos, and larvae is also slowly beginning to emerge as a viable alternative to manual handling and culture (Table 1) (39). Below we will briefly summarize these up-and-coming technologies and address their interdisciplinary promises and future challenges for specific applications.
The nematode Caenorhabditis elegans has recently emerged as a convenient and versatile experimental animal (40). It is the smallest and simplest fully differentiated multicellular organism (featuring only 959 somatic cells in an adult) that can be easily cultivated at very low cost and in large numbers (Table 1). It has a very short life cycle and is particularly suitable for genetic manipulations and analysis. The sequencing of the C. elegans genome in 1998 and later implementation of RNA interference techniques further increased interest in genome-wide screens in this model organism (6, 41). The 60–80% gene homology to human counterparts has stimulated genetic engineering of worms carrying mutations in many highly central biochemical pathways (40, 42, 43). Its short lifespan means that phenotypes can be scored quicker as compared with mammalian models, while transparency allows convenient in vivo imaging and monitoring of cellular processes (Table 1) (6, 40, 44). These features have recently made C. elegans particularly attractive for drug discovery and toxicology (40, 44–46).
C. elegans adult form is approximately 1 mm long and 30 μm wide (Table 1) and therefore can be robotically dispensed into microtiter plates for high-throughput library screening (6). Integrated manipulation, sorting, immobilization, and precise positioning of micron-sized small-model organisms such as the nematodes, however, still represents a challenging task. This is especially true when high-resolution imaging is required to score particular phenotypes or assess effects of drug action at the cellular level. In this context, a plethora of innovative chip-based technologies for cultivation, manipulation, immobilization, and sorting of C. elegans has recently been reported. Below we briefly summarize only several most interesting technologies and classify them according to the particular experimental application.
An elegant solution for a long-term, on-chip cultivation of C. elegans has recently been reported by the Whitesides group at Harvard (47). These authors have described a microfabricated device consisting of an array of 16 micro-chambers holding individual worms (Figs. 3A and 3B). Chambers are individually addressable via a network of microfluidic channels for delivering bacterial nutrition (Figs. 3A and 3B) (47). The microchip technique reportedly enabled observation of many behavioral and physiological phenotypes over the entire lifespans of worms (47).
Figure 3. Chip-based devices for live-animal analysis of C. elegans. (A) Microfluidic chip for life-long observations of nematodes. Design includes an array of 16 chambers and a network of branching channels that deliver a food source (suspension of E. coli). The bypass outlet enables the removal of sedimented bacteria from the inlet. Reproduced by permission of The Royal Society of Chemistry from Ref. (47) (B) Microfluidic chambers with integrated micromechanical worm clamps. Wedge-shaped microchannels are fabricated in soft polymer PDMS. Worm traps have dimensions comparable to the cross-sectional diameter of nematodes. The mechanical immobilization is not damaging to the worms and reversal of the flow can release the worms from the clamps: subsequently cultured, they had typical lifespans and reproduced normally. Note that the width of the microchannel directly to the right of the chamber is just wide enough to allow the passage of a young worm into the chamber. Arrows indicate the direction of fluid flow. Reproduced by permission of The Royal Society of Chemistry from Ref. (47) (C)–(D) Overview of the chip-based, pneumatic nematode trapping system. (C) A 3D virtual prototype showing the bilayer manifold without (left) and with an immobilized worm (right). Note the deflection of the elastomeric membrane that immobilizes the worm in place against the glass cover slip. Reproduced by permission of Nature Publishing Group from Ref. (49). (D) Integrated elastomeric valves (yellow rectangles 1–4) for control of inlet regulation, fine positioning of the worm, and gating to the recovery chambers (upper panel). Cross-sectional profiles of the trap area (lower panel). Note the deflection of the elastomeric membrane increasing with air pressures from 0 to 35, 70, 105, 140, and 175 kPa (left). Trapped worms with membrane deflection at 105 and 140 kPa, respectively (right). Reproduced by permission of Nature Publishing Group from Ref. (49). (E) Integrated chip-based system for automated imaging, phenotyping and sorting of C. elegans. Immobilization of the worms is achieved by gentle vacuum suction. This technology is coupled with customized software to enable a fully automated sample loading, specimen positioning, imaging, and on-the-fly classification of worms based on morphological and intensity features. Reproduced by permission of Nature Publishing Group from Ref. (52). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Trapping and immobilization of live worms without anesthetics can alleviate experimental bias due to potential chemical and physiological interactions of anesthetic agents and considerably simplify the laboratory protocols. Recently several innovative technologies for on-chip immobilization of C. elegans have been reported including integrated micromechanical clamps, pneumatic actuators, microsuction manifolds, and thermo-chemical polymers (48–52).
Hulme and colleagues recently proposed physical immobilization of large numbers of living C. elegans adult forms by using arrays of wedge-shaped microchannels fabricated in soft polymer poly(dimethyl) siloxane (PDMS) (Figs. 3A and 3B). Worm traps have dimensions comparable to the cross-sectional diameter of nematodes (48). The flow of medium automatically distributes individual worms across each clamp and facilitates immobilization of up to 100 worms in less than 15 min (48). Chip-based clamps are reportedly applicable for performing morphological analysis, laser microsurgeries, and high-resolution fluorescence imaging. Importantly, the mechanical immobilization is not damaging to the worms and reversal of the flow can release the worms from the clamps: released worms subsequently cultured had typical lifespans and reproduced normally (Figs. 3A and 3B) (48).
The groups of Chronis and Ben-Yakar have recently pursued an elegant pneumatic actuator approach to immobilize worms on a chip and then interfaced this technology with femtosecond laser nanosurgery (Figs. 3C and 3D) (49). These authors have developed the adaptive deflection of an elastomeric membrane sandwiched between two-layered microfluidic channels that allow the immobilization of the worms from L4 to adult size. Pressurization of the upper channel deflects the thin, elastomeric membrane that immobilizes the worm. This technology holds the worms mechanically immobilized against the glass cover for ideal focusing and precise laser nanosurgery (Figs. 3C and 3D) (49). Importantly, the traps can be adjusted to the size of the biological specimens. Moreover, the chip-based system integrates feeding modules facilitating long-term imaging studies of the worms post-surgery as well as their sorting and screening (49).
Krajniak and Lu have recently also proposed a hybrid microfluidic chip with a thermo-sensitive triblock copolymer Pluronic F127 for repeatable immobilization and long-term imaging of C. elegans at physiological conditions (51). PF127 represents a group of amphiphilic block copolymers that at low temperatures behave like viscous liquids whereas at temperatures above the gelation temperature are capable of non-invasively immobilizing the animals in the gelatinous blocks (51, 53, 54). Importantly, the sol-to-gel transition is reversible allowing lucid animal recovery simply by lowering the temperature of the surrounding medium. The precise regulation of the temperature was achieved by a network of microfluidic heating conduits on the multilayer chip-based device (51). PF127 reportedly had no adverse effects on long-term viability or development of C. elegans. Furthermore, the PF127 solution did not exhibit significant autofluorescence or light scattering, thus enabling precise high-resolution imaging of neurons and synapses along the ventral and dorsal nerve cords of C. elegans larval stages (51).
Sorting and High-Throughput Analysis
LOChip systems encompassing numerous functionalities, robust performance, and automated operation are up and coming to the field of live organism analysis. In this context, a fully integrated high-throughput chip-based system for automated imaging, phenotyping, and sorting of C. elegans has recently been developed by Lu's group at Georgia Institute of Technology (Fig. 3E) (52). The microfluidic device was fabricated using replica moulding in PDMS and features integrated on-chip valves that control a suspension of nematodes. Immobilization of the worms is achieved by a gentle vacuum suction (Fig. 3E) (52). This technology is coupled with customized software to enable a fully automated control of the device including (i) sample loading, (ii) specimen positioning, (iii) imaging, and (iv) on-the-fly classification of worms based on morphological features and fluorescence intensity features (52). The authors have reportedly validated this innovative technology for classification and sorting based on cellular and subcellular phenotypes of C. elegans with over 95% accuracy and speeds reaching several hundred worms per hour (52). Rohde and colleagues have also proposed a similar system for complex whole animal genetic and drug screening routines (55). Their chip-based technology combines various functionalities to provide on a one chip: (i) sorting of live animals; (ii) cultivation in an array of microfluidic chambers; (iii) vacuum immobilization for subcellular-resolution time-lapse imaging without the need for anesthesia, and (iv) interface to provide a multiplexed animal dispenser and large-scale screening of drug libraries (55). The microfabricated sorter can reportedly immobilize and release animals repeatedly in 100 ms intervals. Importantly the device supports three-dimensional cellular and subcellular imaging that cannot be resolved using conventional cytometric systems such as COPAS/BioSorter that feature a flow-through principle and therefore capture only a one dimensional intensity profile of the animal (55).
Interestingly, a droplet-based microfluidic platform where worms can be grown in aqueous microcompartments separated by perfluorocarbon carrier oil has also been proposed by Clausell-Tormos and colleagues (56). Hatched C. elegans worms had been shown to survive and proliferate within the microplugs for several days prospectively enabling high-throughput screening of chemical and genetic libraries (56). Shi et al. have subsequently developed a droplet-based microfluidic system that integrates a T-junction droplet generator together with an innovative droplet trapping array for physical immobilization of encapsulated worms (57). This technology allows for culture of individual worms in a parallel series of segmented flow plugs and provides a highly controllable microenvironment for single-organism behavioral studies (57). These technologies open up new horizons for high-throughput studies and can vastly accelerate the introduction of single-animal resolution bioassays for drug discovery.
D. melanogaster is a traditional metazoan model studied now for well over a century. Fly models have greatly enabled the discovery of key fundamental biological phenomena. They offer a large array of convenient genetic tools, greatly facilitating both forward and reverse genetic approaches (58). Sequencing of the whole D. melanogaster genome in 2000 demonstrated that a high degree of homology exists between invertebrates and humans with over 60% of human genes having a fly counterpart (Table 1) (6, 59). Moreover, the inactivation of homologous Drosophila genes often reflects phenotypes reminiscent of their mammalian counterparts (6). The recent introduction of the whole genome RNA interference (RNAi) screens provided a rapid discovery tool for new putative targets. Fly models are, therefore, increasingly finding noteworthy applications in chemical screening (60). In this context, a drosophila-based whole animal model screen has recently been reported by Das and colleagues to identify therapeutically useful compounds against thyroid cancer (61). These findings validate the use of fly models for both high-throughput drug discovery as well as identification of beneficial drug combinations (10, 61).
Conventional manipulation and analysis of D. melanogaster eggs and embryos is performed manually. Some advances in liquid handling robots can nowadays considerably automate dispensing of fly embryos for large-scale genetic screening but suffer from pipetting errors, evaporative medium loss, and considerable complexity. When imaged in conventional vessels, embryos are mostly oriented along their major axis parallel to the coverslip or well. Owing to the embryo small size and thus difficult mechanical manipulation, this essentially precludes observations of dorsoventral signal propagation in response to drugs (62). The size of fly eggs and embryos (approximately 100 μm) make them, however, perfectly suitable for integrated chip-based systems that can provide a superior degree of automation and environmental control (Table 1). Microfluidic systems can also provide innovative ways to precisely control animal position for imaging. Recently, an approach based on passive hydrodynamics, has been developed by Chung and colleagues to rapidly trap and position hundreds of Drosophila embryos on a microfluidic chip (62). The device, fabricated in polydimethylsiloxane (PDMS), consists of a serpentine 700 μm wide fluid-delivery manifold for robust handling of nonspherical objects and an array of approximately 700 cross-flow channels (Figs. 4A–4C) (62). Each cross-flow channel includes a shortened tubular trap. The shape of the traps allows for upright embryo positioning (Figs. 4A–4C). Interestingly, the trapping mechanism does not rely on a fluid resistance change following the occupation of traps and as such allows for very dense arraying of embryos with nearly 90% trapping efficiency (62). This innovative design reportedly enables high-throughput and quantitative analysis of multiple morphogen gradients in the dorsoventral patterning system (62).
Figure 4. Microfluidic technologies for the analysis of D. melanogaster. (A)–(C) A high-density microfluidic embryo-trapping array. (A) Design characteristics of the embryo-trap technology. The device is fabricated in PDMS and consists of a serpentine 700 μm wide fluid-delivery manifold for robust handling of nonspherical objects and an array of approximately 700 cross-flow channels. Each cross-flow channel includes a shortened tubular trap. The shape of the traps allows for hydrodynamic embryo immobilization and vertical positioning. Scale bar: 500 μm. Reproduced by permission of Nature Publishing Group from Ref. (62) (B) Scanning electron micrograph of the single embryo-trap structure. Scale bar: 100 μm. Reproduced by permission of Nature Publishing Group from Ref. (62) (C) Steps of the embryo trapping and immobilization: (i) embryo is directed into the trap (left), (ii) the fluid flow around the embryo orients it vertically (middle), and (iii) microfabricated trap contracts and secures the embryo (right). Imaging focal plane—yellow. Direction of the main fluid flow—blue arrows. Direction of the secondary stream hydrodynamically immobilizing the embryo—red arrow. Reproduced by permission of Nature Publishing Group from Ref. (62) (D) Spatial and temporal stimulation of a live Drosophila embryo by using two laminar streams flowing in a ‘Y’ junction microfluidic device made in biologically compatible polydimethylsiloxane polymer. Note that laminar flow was used to create two distinct flows of warm and cold buffer around Drosophila embryo suspended in the cross-section of the channel. Reproduced by permission of The Royal Society of Chemistry from Ref. (64) (E) Analysis of the temperature profile around a live embryo in a microfluidic device. Note that the temperature profile was analyzed in real-time using a suspension of thermochromic liquid crystals and a tight thermal boundary is clearly observed between the two laminar streams flowing around the Drosophila embryo. Reproduced by permission of The Royal Society of Chemistry from Ref. (64). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Recently, the group of Ismagilov has also shown spatial and temporal stimulation of live Drosophila embryos by using two laminar streams flowing in a simple “Y” junction device made in polydimethylsiloxane (Figs. 4D and 4E) (63, 64). Microfluidic laminar flow was used to create two distinct flows of warm and cold buffer around a Drosophila embryo suspended in the cross-section of the channel (Figs. 4D and 4E). This was greatly facilitated by the low thermal conductivity of the PDMS. In this elegant work, the authors characterized the effects of positioning of the embryo in the channel on fluid flow and the temperature distribution around the embryo (63, 64). They also showed for the first time how variations in the bicoid morphogen gradient can compensate the effects of extremely unnatural environmental conditions, where the two extreme halves of the embryo are developing under drastically different temperatures (63). Dagani and colleagues have developed a further automation of this unique technology by using a self-assembly technique for immobilizing embryos on an open substrate under a minimal potential-energy principle (65). Similar microfluidic techniques could prove very useful for spatio-temporal control of local environmental changes and warrant previously unattainable studies on how biochemical networks respond to perturbations by both fluctuations in environmental conditions and genetic variation (63, 64).
Yet another interesting work by Zappe and colleagues has recently presented development of an automated microelectromechanical (MEMS) system for large scale microinjections to Drosophila embryos (66). Innovative hollow needles were made of silicon nitride. These were used to deliver on average 60 pl RNAi solution per embryo from an 500 nl integrated on-chip reservoir (66). Volume regulation is provided by an off-chip air pressure pulse, precisely dosed by a microcontroller. Injections can be reportedly delivered within tens of milliseconds dramatically improving throughput of gene delivery. Moreover, the chip-based microinjection system can be operated in parallel allowing analysis of more than 50 genes per day (66). As claimed by the authors, this reflects an improvement of more than ten times compared to manual procedures. Application of this technology for high-throughput RNA interference (RNAi) screens will likely accelerate whole genome screens and improve efficiency, accuracy, reliability, and controllability of RNAi experiments on fly embryos (66).
An interesting attempt has also been made to develop a dedicated D.melanogaster sorter that integrates robotics, flow cytometric principles, and microcapillary fluidics (67). The sorter comprises a 400 μm square glass capillary embedded in a holder with immersion oil. A peristaltic pump drives the solution containing D.melanogaster embryos into the capillary at a fixed rate of 6 ml/min (67). The capillary is designed to restrict the embryos to only two possible orientations, thus facilitating rapid flow cytometric interrogation by a 488 argon ion laser (67). The sorting mechanism is an electro-mechanical switch positioned between two thin walled fluidic lines. It comprises a rare-earth neodymium supermagnet driven by two electromagnets. Actuation of the opposite electromagnets exerts mechanical force that moves the magnet, sequentially opening the collection or waste fluidic manifold (67). According to the authors, the electro-mechanical switch can actuate at a speed of 10 ms. This gives a theoretical sorting speed of approximately 100 D.melanogaster embryos per second (67).
Finally, Stelzer et al. have recently developed an innovative imaging technique referred to as selective plane illumination microscopy (SPIM) that allows to generate real-time and multidimensional images of large biological specimens (68). This noteworthy technology is particularly suitable for the lifelong cytometric analysis of small model organisms and opaque embryos. Optical sectioning of samples during SPIM imaging is achieved by illuminating the biological sample along a separate optical path orthogonal to the detection axis (68). Interestingly, only the plane under observation is illuminated and this considerably reduces photo bleaching and photo toxicity effects (68). As a result, the total number of fluorophore excitations required to image a 3D sample is significantly reduced compared to conventional laser scanning microscopy (LSM). SPIM has already been applied to real-time analysis of developing Drosophila embryos embedded in low-melting point agarose cylinders. Real-time imaging for up to 3 days did not induce any noticeable effects on embryogenesis and development (68). The SPIM method is therefore non-disruptive and easily applicable to living embryos. Although it has not been interfaced with chip-based devices as yet it provides a unique set of lifelong imaging capabilities that could be explored further using innovative microfluidic technologies.
Zebrafish (Danio rerio) is receiving increasing attention as a genetic model of human disease and platform for accelerated drug discovery (8, 15). Simple and cost-effective maintenance together with abundant experimental techniques and molecular tools have made zebrafish the model of choice for chemical genetics and large-scale in vivo drug screening routines (7, 8, 69, 70). The zebrafish has a short reproductive cycle that is coupled with a large number of progeny and relatively small amount of space needed to maintain large culture of offspring at a reasonably low cost (16, 70–73). Small size, optical transparency of organs, and easy of culture make zebrafish embryos, larvae, and juveniles the ideal model for large scale genetic and pharmacological studies (Table 1) (7, 8). Several zebrafish models of human diseases have already been developed where physiological processes can be rapidly assayed using sophisticated 4D microscopy in the native context of the developing organism (74, 75). All these characteristics have led to the realization that lead pharmaceutical discovery, toxicology screening, and regenerative medicine studies can be undertaken using D. rerio owing to the high physiological conservation to humans and applicability to perform chemical genetic screens (7, 8, 13, 15, 70, 72). It has been also recently proposed that transgenic zebrafish could be designed to rapidly detect low levels of chemical contaminants such as heavy metals or pesticides using pollution-inducible response elements (16, 71, 73, 76). Such systems provide very sensitive, economical, and environmentally relevant biomonitoring solutions applicable for automation and potential field deployment (71, 77).
Despite some progress in large object sorting as discussed above, the embryo and larvae handling, sorting and treatment is, still predominantly performed manually under static microtiter plate-based conditions. This limits research productivity. The development of integrated Lab-on-a-Chip technologies for automated manipulation of D. rerio is still in its infancy but can prospectively accelerate drug discovery pipelines. One of the earliest attempts to provide on chip manipulation and analysis of Danio rerio embryos was the application of segmented flow for toxicological and drug screening studies (Figs. 5A and 5B) (78). Zebrafish embryos were successfully manipulated using segmented flow with perfluoromethyldecalin (PP9) as the carrier liquid inside the Teflon (PTFE) tubes. The single liquid phase embryo suspension was separated through the carrier liquid using the immiscibility of the phases. Aqueous microsegments containing single embryos were approximately 0.5 cm in length with a volume of approximately of 6 μl (Fig. 5B). Fish development in microsegments was reportedly unaffected over a time period of up to 80 hours (78). Interestingly, a relative humidity of 80% within the incubator was sufficient to limit the evaporative loss of the PP9 solvent (Figs. 5A and 5B). Moreover, oxygen diffusing through the tubing and PP9 was sufficient to support the growth of the embryos in the aqueous microsegments (78).
Figure 5. Fish-on-chips—microfluidic technologies for the analysis of zebrafish. (A)–(B) A micro-segment flow technology for screening and development studies. (A) Experimental setup. Note that a computer-controlled syringe pump was used for generating segmented flow with perfluoromethyldecalin (PP9) as the carrier liquid inside the Teflon (PTFE) tube coil. The single liquid phase embryo suspension was separated through the carrier liquid using the immiscibility of the phases. Aqueous microsegments containing single embryos were approximately 0.5 cm in size with a volume of approximately of 6 μl. Reproduced by permission of The Royal Society of Chemistry from Ref. (78). (B) Multicell stage of D. rerio embryo inside an aqueous microsegment embedded by PP9 carrier liquid in a Teflon tube. Reproduced by permission of The Royal Society of Chemistry from Ref. (78). (C)–(D) Transport of live zebrafish embryos using digital microfluidic EWOD technology. (C) Principles of EWOD technique in which a droplet of aqueous solution is confined by two plates and the dielectric layer with a hydrophobic surface coating: t, thickness of the dielectric; d, gap spacing; q and q0, advancing and receding contact angles, respectively; Device was able to transport an 0.5 mm diameter zebrafish embryo in 20 μl droplet of E3 medium. Reproduced by permission of The Royal Society of Chemistry from Ref. (79). (D) Manipulation and transport of zebrafish embryo using 20 μl droplets and EWOD droplets. Reproduced by permission of The Royal Society of Chemistry from Ref. (79). (E) The vertebrate automated screening technology (VAST) for high-throughput manipulation and 3D imaging of zebrafish larvae at cellular resolution. VAST comprises robotics and microcapillary fluidics to provide multiple step operation such as: loading, detection, positioning, rotation, focusing, imaging, laser manipulation, and subsequent dispensing of D. rerio larvae. Inset shows imaging of a transgenic, fluorescent zebrafish larva. For detailed description refer to text. Reproduced by permission of Nature Publishing Group from Ref. (9). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Electrowetting-on-dielectric (EWOD) technique for transport of live zebrafish embryos has recently been presented by Son and colleagues (Figs. 5C and 5D) (79). Droplet microfluidics involves manipulating discrete liquid samples sandwiched between two plates (80, 81). The bottom plate usually includes microelectrodes hidden beneath a dielectric layer while another plate serves as a common ground electrode (Fig. 5C) (79). Droplet movement can be freely controlled in two dimensions by applying an electrical potential across microelectrodes (79, 81). This creates an electromechanical force manifested as EWOD (Fig. 5C). Son and colleagues have demonstrated the feasibility of automatically addressable electrodes and digitally programmed transport of zebrafish embryo within droplets in a two-plate digital microfluidic device (Fig. 5D) (79). Despite some electrolysis and Joule heating, embryos transported in droplets for up to two hours remained viable and developed normally. Interestingly, these authors have for the first time demonstrated the applicability of the EWOD technique for dechorionation of D. rerio embryos by mixing a droplet of digestive reagent (Pronase) with a droplet containing the specimen (79).
Another integrated technology for in vivo chemical and genetic screening on zebrafish larvae has recently been reported by Yanik's group at MIT (9). The vertebrate automated screening technology (VAST) allows for automated manipulation and imaging of zebrafish larvae at cellular resolution in three dimensions (Fig. 5E) (9). This innovative technology comprises robotics and microcapillary fluidics to provide multiple step operation such as: loading, detection, positioning, rotation, focusing, imaging, laser manipulation, and subsequent dispensing of D. rerio larvae (Fig. 5E) (9). Larvae are detected using a photodiode and two LEDs where a simultaneous monitoring of the transmitted and scattered light precisely discriminates biological specimens from residual air bubbles and/or debris (Fig. 5E). The reliability of this innovative detection system is reportedly nearly 100%. Following detection, larvae are loaded into the microcapillary using a stepper motor-driven syringe pump and placed under a high-resolution imaging system. Another innovation of VAST includes a 3D-axis computer-controlled stage that can finely manipulate and position the assembly. The capillary can be also rotated along the longtitudal axis (Fig. 5E). This facilitates not only subcellular imaging of organs from multiple angles but also in vivo optical manipulations such as femtosecond laser microsurgery and localized activation of fluorescent reporter probes (9).
Recently yet another unique robotic system that combines mesofluidic circuitry has been developed by Knapp's group at the Centre Suisse d'Electronique et de Microtechnique (CSEM SA, Alpnach, Switzerland). ZebraFactor is reportedly capable of sorting and dispensing individual embryos in a highly reproducible manner (Fig. 6). Sort decisions are performed using a fast imaging system combined with an on-the-fly image analysis algorithm which is capable of identifying a set of predefined optical characteristics (39). Embryos can be then automatically be extracted from the sorter and dispensed to a multiwell plate. ZebraFactor operates with an average speed of 8 seconds per single embryo (Fig. 6) (39). This corresponds to loading a 96-well multi-titer plate in about 11 min which is comparable to manual dispensing performed by skilful technical personnel. The robotic system can, however, operate in a continuous manner, achieving overall higher throughput per day while at the same time maintaining high levels of reproducibility. The survival rate of sorted zebrafish eggs was reportedly above 90% (39).
Figure 6. XenoFactor and ZebraFactor technology image-based fluidic sorting systems for automated sorting and microinjection of zebrafish and Xenopus. (A) Device overview consisting of the CellSorter, Microinjection unit, and WellPlateFeeder. (B) CAD design of the sorting unit with a two-camera configuration to sort opaque Xenopus oocytes. (C) Principles of the fluid and object motion using the sliding ring technology incorporated in the sorting unit. The eggs/embryos are driven by drag and friction forces and rolls along the base of the sliding ring. Data courtesy of CSEM SA, Alpnach, Switzerland (www.csem.ch) (39). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Delivery of two-dimensional patterns of tracer molecules, DNA, and mRNA into living zebrafish embryos has been achieved using a combination of electroporation and microfluidics (82). For this purpose, Bansal and colleagues have used platinum electrodes, microfabricated into various shapes and passivated with silicone elastomer. Single square 50–100 ms pulses of 0.20–0.40 kV/cm were used to create transient pores and introduce compounds into the developing embryos (82). This approach allows for an innovative way to modify spatial and temporal pattering genes, proteins, and bioactive molecules. It can be particularly attractive in studies of embryonic development and morphogenesis where heterogeneous spatial regulation of developmental programs is difficult to control with existing experimental tools (82).
Apart from laboratory automation in handling the specimens, another major factor limiting the throughput of zebrafish chemical screens is still lack of dedicated software algorithms to automatically detect and quantify specific structures within the living organism (83, 84). The complexity of images acquired on multicellular organisms such a zebrafish make detecting and classifying small biological changes difficult (83, 84). Therefore, image data analysis is commonly performed manually throughout the screening procedure. This considerably restricts throughput and introduces large experimental bias. Some progress has recently been made by Vogt and colleagues, who have described a system for automated imaging and analysis of zebrafish embryos (83, 84). These authors have provided evidence that image capture on an ImageXpress Ultra laser scanning confocal reader coupled with custom developed image analysis algorithms based on Definiens Cognition Network Technology can automatically quantify GFP expression in the heads of transgenic embryo with a high level of accuracy and reproducibility (83, 84). Another interesting recent work has shown automatic detection and quantification of pigments in zebrafish embryos (85). The algorithm can identify the head vs. torso and then the boundaries matching the back and abdomen of zebrafish embryos (85). Further development of real-time cytometric algorithms and standardized bioassays for the analysis of multicellular specimens is, however, required to achieve throughputs matching the needs of industrial drug discovery.
Amphibians such as African clawed frogs have contributed to a rapid progress in developmental genetics and cell and regenerative biology during the last two centuries (86). They offer comparable experimental advantages that have favored the usage of zebrafish models, including extrauterine development, transparency of developing eggs and larvae, and permeability for small molecule drugs (Table 1) (15, 86). Xenopus oocytes, fertilized eggs, embryos, and tadpoles are also often used in conjunction with various techniques such as microsurgery, mRNA injections, gene knockdown studies, and real-time video-microscopy among others (15, 86). Amphibians as a tetrapod are also evolutionarily nearer to humans than zebrafish with more conservative diploid genome structure. Moreover, their internal organs better reflect morphological and physiological functionalities of their human counterparts than those of, e.g., zebrafish. In this context, recent noteworthy reports suggest that both Xenopus embryos and tadpoles can provide a viable alternative to zebrafish forchemical and whole-organism drug discovery screens (15, 87, 88). Moreover, Xenopus can be a highly cost effective and robust model for estimation of drug toxicity and health and environmental hazards (15, 17, 88, 89) as evidenced by the Frog Embryo Teratogenesis Assay-Xenopus (FETAX) test for 96-h whole-embryo developmental toxicity screening (17, 89–91). The scope of Xenopus applications in modern biomedicine and toxicology necessitates the need for automated, in situ, miniaturized systems for high-throughput handling and analysis of amphibian oocytes, embryos, and larvae.
Conventional in situ manipulation and analysis of tadpoles reportedly involves standard multiwell plate systems and time consuming, manual placement of individual animals (92). Moreover, immobilization of tadpoles during imaging requires anesthetization. This is labor-intensive and introduces additional experimental bias due to potential chemical and physiological interactions of anesthetic agents (92). To overcome these limitations a very interesting technology for rapid analysis of transgenic and fluorescent Xenopus tadpoles has recently been proposed by Fini and colleagues. They demonstrated for the first time a flow-through fountain flow cytometry (FFC), originally developed for real-time detection of bacteria and protozoa in aquatic samples (92–94). The FFC technology developed by Finni et al. is a real-time system where tadpoles flow though a flow cell towards an interrogation window integrated with a digital camera and blue (488 nm) LED excitation source. Gravity-driven flow was used to continuously circulate tadpoles through the flow cell between two reservoirs (92). The technology was validated using transgenic Xenopus laevis tadpoles harboring a chimeric gene with a heavy metal responsive element fused to a green fluorescent protein (metallothionein promoter from zebrafish; MTZF-eGFP) (92). This transgene can be selectively induced by the presence of heavy metals resulting in a convenient readout in the form of a bright green fluorescence of the whole animal. This work provides an interesting adaptation of the miniaturized, continuous flow techniques for rapid imaging of small aquatic animals. Comparative analysis has shown a high level of sensitivity as compared to conventional static imaging while real-time analysis greatly accelerates the data acquisition and supports implementation of transgenic Xenopus models for heavy metal biomonitoring (92). Importantly, the FFC technology permits for a non-invasive and real-time examination of Xenopus larvae without any anesthetic agents or extensive mechanically manipulations. This greatly minimizes stress and yields statistically reliable data (92).
Another interesting example is the XenoFactor technology developed at CSEM in Switzerland and specifically designed to work with cells in the millimeter range such as Xenopus laevis oocytes (Fig. 6). It combines microfluidics with robotics to provide fully automated processing of hundreds of Xenopus oocytes. CSEM XenoFactor utilizes similar design to CSEM ZebraFactor technology described above (Fig. 6) (39). Sedimentation of large objects is prevented by constantly rotating them in a circular channel called a storage ring (39). Motion of oocytes is facilitated by a sliding ground and fixed walls and cells can be focused in front of the CCD imaging system to ensure that only a single cell passes though the interrogation and sorting point (Fig. 6). The optical system consists of a CCD recording 100 frames per second and used for observing both transparent and opaque cells (39). Object parameters such as size, shape, and transparency can be analyzed by specifically designed on-the-fly imaging algorithms. Reportedly large numbers of Xenopus oocytes can stored in the rotating storage ring (39). Single oocytes can be further sorted based on the preselected parameters and the system gives output speeds of up to one oocyte every four seconds. This is a remarkable 32-fold increase of speed as compared with conventional methods. Interestingly, XenoFactor has been reportedly interfaced with an integrated microinjection system (Fig. 6). An integrated system is capable of automatic sorting, immobilization, and microinjection of oocytes that are subsequently re-analyzed and stored in a separate container for subsequent processing (39).
Some progress is also being made in the development of an integrated system for electrophysiological measurements on Xenopus oocytes. In this context, Dahan and colleagues have recently developed a non-invasive chip-based technique that replaces the traditional time consuming and invasive “two-electrode voltage-clamp” (TEVC) method (95). Reportedly, the technology couples non-invasive voltage-clamp measurements with rapid fluidic exchange. Oocyte immobilization is performed by positioning of non-devitellinized oocytes on an aperture on top of the perfusion microchannel (95). Reagent solutions are subsequently delivered to the chip by two computer controlled syringe pumps. Fluidic exchange has been reduced to 20 ms, providing new vistas for performing complex pharmacological protocols and making it suitable for screening of ion channel ligand libraries on live Xenopus oocytes (95).