Wormometry-on-a-chip: Innovative technologies for in situ analysis of small multicellular organisms

Abstract

Small multicellular organisms such as nematodes, fruit flies, clawed frogs, and zebrafish are emerging models for an increasing number of biomedical and environmental studies. They offer substantial advantages over cell lines and isolated tissues, providing analysis under normal physiological milieu of the whole organism. Many bioassays performed on these alternative animal models mirror with a high level of accuracy those performed on inherently low-throughput, costly, and ethically controversial mammalian models of human disease. Analysis of small model organisms in a high-throughput and high-content manner is, however, still a challenging task not easily susceptible to laboratory automation. In this context, recent advances in photonics, electronics, as well as material sciences have facilitated the emergence of miniaturized bioanalytical systems collectively known as Lab-on-a-Chip (LOC). These technologies combine micro- and nanoscale sciences, allowing the application of laminar fluid flow at ultralow volumes in spatially confined chip-based circuitry. LOC technologies are particularly advantageous for the development of a wide array of automated functionalities. The present work outlines the development of innovative miniaturized chip-based devices for the in situ analysis of small model organisms. We also introduce a new term “wormometry” to collectively distinguish these up-and-coming chip-based technologies that go far beyond the conventional meaning of the term “cytometry.” © 2011 International Society for Advancement of Cytometry

Cytometry has traditionally been associated with multivariate studies of single cells and distinctive cell subpopulations (1, 2). Speed and multiparameter data collection have significantly contributed to the dramatic expansion and popularity of cytometric techniques in cell biology (Table 1) (3). Interestingly, cytometric and photonic technologies have recently begun to find noteworthy applications in analysis of small model organisms (4). Invertebrate (mainly the nematode Caenorhabditis elegans and the fruit fly Drosophila melanogaster) and small vertebrate animal models (mainly the clawed African frog Xenopus laevis and zebrafish Danio rerio) are gaining considerable interest in drug discovery and toxicology (5–10). They offer substantial advantages over cell lines and isolated tissues, providing analysis of cells in the context of cell–cell and cell–extracellular matrix interactions and under normal physiological milieu of the whole organism (8, 11, 12). This provides analytical capabilities that cannot be easily replicated in vitro, like organogenesis, tissue regeneration, drug accumulation, metabolism and organ-specific toxicity. Small size, optical transparency of organs, and ease of husbandry make small model organisms convenient for large scale genetic and pharmacological studies (Table 1) (9, 13). Compound library screens can also be performed with a remarkable throughput as compared to, e.g., rodent models (6–8, 13, 14). These can be used to find new chemical structures and putative drug targets where libraries are tested on wild-type animals for their toxicity and/or emergence of specific phenotypes (10). Alternatively and perhaps more excitingly, genetically modified models can be designed using “knock down” (loss-of-function diseases) or “knock in” (gain-of-function diseases) techniques to reproduce relevant human disorders (9, 10). Undoubtedly the physiological differences between humans and invertebrates and lower vertebrates are significant and as such drug discovery approaches using these models can often yield an incomplete picture of the human symptoms. Despite these limitations, however, small multicellular animal models are important tools to bridge the gap between traditional in vitro and inherently costly and ethically controversial rodent animal assays (6, 10, 15). Moreover, a large number of unique environmental and ecotoxicological tests can be performed by using small multicellular animal models (16, 17). This dramatically expands the potential scope for the development of in situ cytometric technologies.

Table 1. Characteristics and applicability of small multicellular animal models for an automated handling and cultivation on miniaturized lab-on-a-chip devices

Model features	Invertebrates		Vertebrates
	C. elegans	Drosophila	Zebrafish	Xenopus	Chicken	Mouse
Evolutionary similarity to humans	Very distant	Very distant	Distant	Intermediate	Intermediate	Close
Generation time	3–5 days	10–14 days	3–4 months	4–6 months	20 weeks	3–4 weeks
Average offspring size	300	90	150	2500	1	7
Average embryo size	50 μm	100 μm	1.2 mm	1 mm	4 cm	8 mm
Average adult size	1 mm	3 mm	6 cm	10 cm	40 cm	10 cm
Cost per assay	Low	Low	Low	Low	High	High
Screening throughput	High	High	Medium	Medium	Low	Low
In situ automation applicability	+++	+++	+++	+++	−	−
LOC applicability	+++	+++	+++	+++	−	−
Current advancement of LOC technologies	+++	+	+/−	+/−	−	−

Notwithstanding this, cytometric in situ analysis is still deeply in its infancy. This is mostly due to the inherent limitations of conventional techniques, such as flow cytometry, that make dynamic analysis of large metazoan organisms difficult. Although, as we describe in this article, some fascinating progress has recently been made in the field of large particle analysis and cell sorting, many desirable functionalities for robust and automated in situ analysis are still absent. These include integrated (i) specimen loading, (ii) specimen trapping and immobilization, (iii) repeated time lapse data acquisition, (iv) real-time stimulation/perfusion without mechanical disturbance, (v) sorting, (vi) post-analysis recovery, and (vii) extended culture without human intervention. In the context of advanced functionalities, the last decade in particular has brought many spectacular innovations to the field of cytometric technologies. Rapid progress in physics, electronics, as well as material sciences has facilitated the development of miniaturized bioanalytical systems collectively known as Lab-on-a-Chip (LOC). LOC represents the next generation of analytical laboratories that have been miniaturized to the size of a matchbox and represent one of the most groundbreaking offshoots of nanotechnology and microelectronics.

Microfluidic technologies combine micro- and nanoscale science, allowing the application of laminar fluid flow at ultralow volumes in the spatially confined chip-based micro-channel circuitry. LOC technologies are examples of the most innovative and cost-effective approaches toward the advancement of cytometry and promise new functionalities that can overcome current limitations while at the same time promising greatly reduced costs, increased sensitivity, and ultra-high-throughputs. LOC has already been widely heralded as an emerging technology with a multitude of applications in cell biology. Global market for these technologies has been recently estimated at six billion US dollars. Not surprisingly, implementation of innovative miniaturized systems in integrated culture and analysis of small model organisms is attracting a rapidly growing interest in the drug discovery community (Table 1). In this work we provide a snapshot of the most fascinating technologies that open up new vistas for many fields of modern medicine, drug discovery, biotechnology, and environmental biomonitoring.

Conventional In-Situ Cytometry

A gold standard for in-flow analysis and sorting of small multicellular organisms such as C. elegans, D. melanogaster, Daphnia, marine larvae, Medaka, mosquitoes, sea urchins, Xenopus, zebrafish, and zooplankton has been developed by Union Biometrica Inc. (Holliston, MA, USA) (4, 18). The Complex Object Parametric Analyzer and Sorter (COPAS) and BioSorter Platforms (Union Biometrica) reflect standard flow cytometric principles and are capable of analyzing, sorting, and dispensing objects ranging in size from 250 up to 2,000 μm (Fig 1) (4, 18). Flow cytometric analysis is performed using an axial light-loss detector measuring time-of-flight (TOF) while optical density (extinction, EXT) is determined by the total integrated signal of the blocked light (4, 18). TOF and EXT parameters are indicators of the length and size/internal structure of the multicellular organism, respectively. The fluorescence intensity can be simultaneously measured with TOF and EXT parameters at three different wavelengths using up to three separate fiber optic laser excitation sources. Furthermore, the COPAS can sort and dispense objects based on preselected optical characteristics into microtiter plates or other bulk vessels. For this purpose, conventional cell sorters mostly use electrostatic charge or mechanical/piezoelectric sorting principles that can be potentially harmful to multicellular organisms. To overcome these limitations, the COPAS sorter employs an innovative sorting design to recover unharmed and viable specimens. A pneumatic manifold located after the large bore flow cell removes particles not meeting the predefined sort criteria (deemed negative) by short bursts of compressed air (Fig. 1). While positive specimens are deposited in droplets of medium into the primary container, the negative samples are moved to a secondary vessel (4, 18). The sorting accuracy of the instrument has recently been estimated at 99.4% with sorting purity at 98.8% when approximately 3,097 zebrafish eggs were sorted into 96-well microtiter plates. At the same time 94.7% of the sorted embryos remained viable (Fig. 1) (4). Interestingly, the COPAS system has also been successfully used for dispensing of Daphnia magna for the acute mobilization test showing no effects on the viability of D. magna while bringing a substantial level of automation to this standard OECD ecotoxicological bioassay (Union Biometrica). Despite substantial advantages and efficient automation, the COPAS system still suffers from limitations inherent to conventional flow cytometry where organisms are suspended in a laminar stream of fluid and only integrated signals are collected when an object passes the laser interrogation point (1). A remarkable improvement would be application of a fast in-flow imaging technology similar to that recently mainstreamed as the Image Stream X System (Amnis Corp, Seattle, WA, USA) (19, 20). Development of in-flow imaging cytometry has been pioneered by Kachel et al. in 1979 to provide an innovative method of morphological analysis of cells (21). Gathering morphological information by transmission imaging of cells in flow reportedly requires much shorter exposure times than conventional video microscopy (21). The original instrument was capable of acquiring up to 150 pictures per second and could work as a flow microscope where cell pictures were stored on the 16-mm film (21). The contemporary systems pioneered by Morrissey's group at Amnis Corp (Seattle, WA, USA) integrates a fast in-flow digital imaging with the conventional layout of the flow cytometer (20, 22). At its core the time-delay-integration (TDI) technology preserves sensitivity and image quality during imaging of fast moving cells. In combination with a fast charged couple device (CCD), instead of PMTs, it is capable of simultaneously acquiring six spectrally decomposed images from each cell (20, 22, 23). By combining attributes of both flow cytometry, morphological and fluorescent image analysis, the multispectral in-flow imaging cytometry appears to be a very attractive instrumentation for multiparameter analysis of small model organisms.

Figure 1. Live-animal flow cytometry. (A) Fluorescently activated, large object BioSorter (Union Biometrica) reflects standard flow cytometric principles and is capable of analyzing, sorting, and dispensing objects ranging in size from 250 up to 2,000 μm. (B) Overview of the sorter design. (C) Optical path of the BioSorter. (D) Fluorescent image of D. rerio larva and the corresponding intensity profile. Profiler II software digitizes the object into a succession of peaks and valleys that directly trace the optical density and fluorescence intensity of the object as it passes through the BioSorter flow cell. Profiling enables sorting based on the peak height, peak width, number, and location of peaks within the larvae. Data courtesy of Union Biometrica, Holliston, MA, USA (www.unionbio.com). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

Apart from development of dedicated large particle cell sorters, some noteworthy reports attempted to modify standard electrostatic cell sorters for analysis and sorting of small model organisms. In this regard, Stoeckius and colleagues adapted a FACSVantage SE (BD Biosciences) for generation of synchronized C. elegans embryos expressing OMA-1-GFP fusion protein under the control of oma-1 promoter (24). For this purpose the cell sorter was equipped with a 100 μm nozzle, operating at a nominal pressure of 8 psi and 14,600 Hz frequency. Sorting speeds reportedly averaged at 400–500 embryos per second. The embryoFACS (eFACS) technique was found sufficient to yield tens of thousands of 1-cell stage embryos at a purity of approximately 70% (24). Interestingly, Fernandez et al. have recently modified a BD FACSAria sorter (BD Biosciences) to sort the 250 μm long first larval stages (L1) of C. elegans (25). The authors have used a 100-μm nozzle with sheath fluid pressure at 20 psi and the deflection plate voltage at 6,000 V. Drop-drive frequency was also reduced down to ∼16.4 kHz reducing the potential damage to the worms (25). Interestingly, sorting using this modified live-animal FACS (laFACS) technique had no impact on larvae viability and permitted for a rapid analysis of large quantities of live genotyped worms from a mixed population using GFP-marked balancer chromosomes (25). Sort speeds of up to 800 worms per second were reportedly achievable (25).

Slide based cytometers that combine advantages of both flow cytometry (FCM) and fluorescence image analysis (FIA) can also find noteworthy applications in high throughput analysis of multicellular organisms. In this context, a very interesting technology has recently been developed by Trophos SA (Marseilles, France) under the trademark Plate RUNNER HD® (previously also advertised as the Flash Cytometer®) (Fig. 2). This high-speed imaging cytometer was custom designed to meet the challenges of HTS compound library screening routines. A unique design allows for a large field of view (8 mm) that covers the entire surface area of a single well on a 96-well microtiter plate. Plate RUNNER HD features a maximal optical resolution of 1 μm and depth of field of about 8.5 μm (Fig. 2). Moreover, the multiparameter image acquisition is supported by up to three wavelength excitation/emission optics and on-the-fly digital image analysis. A high-content Accumen ^eX3 microplate laser scanning cytometer (TTP LabTech, Melbourn, UK) has also been applied for C.elegans phenotyping. This innovative technology employs cytometric principles rather than conventional image analysis to collect high-content data. A multiple laser configuration and up to 12 detection channels offer a noteworthy advancement towards a fast and user-friendly approach to high content scanning cytometry (1, 26). It remains to be seen whether other innovative scanning and imaging cytometers such as the LEAP™ and Celigo™ systems (Cyntellect Inc., San Diego, CA, USA), the Opera™ High Content Screening system (Perkin-Elmer, Waltham, MA, USA), InCell Analyzer 1000™ (GE Healthcare Biosciences, Pittsburgh, PA, USA), and the BD Pathway™ 435 and BD Pathway™ 855 analyzers (BD Biosciences, San Jose, CA, USA) will find a new spectrum of possibilities for the automated analysis of small model organisms (1).

Figure 2. High-speed imaging cytometry for live-animal analysis. (A) Plate RUNNER HD® (Trophos SA), a custom designed technology to meet challenges of high-throughput compound library screening routines on C. elegans and zebrafish embryos. (B) and (C) C. elegans and zebrafish fluorescent images, respectively. Data was acquired during large field of view scanning mode that covers the entire surface area of a single well on a 96-well microtiter plate. Multiparameter image acquisition is supported by up to three wavelengths excitation/emission optics and on-the-fly digital image analysis. Data courtesy of Trophos SA, Marseille Cedex, France (www.trophos.com). [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

Miniaturized In-Situ Cytometry

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 (R_e), 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.

Caenorhabditis elegans

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.

Lifelong Cultivation

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.]

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.

Drosophila melanogaster

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.]

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

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.]

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.]

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.

Xenopus laevis

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).

Conclusions

High throughput and content rich automated analysis of small model organisms is still a challenging task. Manual handling procedures are not only time consuming but also error prone, limiting reproducibility and introduction of industry standards. Moreover, the profound lack of in situ technologies that combine automated positioning, sorting, as well as pharmacological, mechanical, and genetic manipulations, and real-time analysis remains the key obstacle to high-throughput organism-based phenotypic assays in drug discovery.

Multidisciplinary advances in integrated microelectromechanical systems (MEMS) are bringing an increasing number of sophisticated technologies that fulfill the promise of real “Lab-on-a-Chip.” Microfabricated devices are now being widely considered as an enabling technology in cell biology, drug discovery, and point-of-care diagnostics (30, 35). Based on the accelerating progress in the field, we envisage that application of microfabricated technologies will also be greatly beneficial in studies on small model organisms. They will offer numerous advantages currently inaccessible for pharmacology and developmental and regenerative medicine. There is a strong reason to believe that recently introduced whole organism phenotypic screens, drug screening routines, laser nanosurgeries, in vitro fertilization, and ecotoxicology testing can be automated, to a previously unanticipated level, by using innovative chip-based in situ devices.

Apart from drug discovery, fascinating prospects are emerging for development of integrated chip-based systems that can greatly support exobiology. Small satellites (nanosatellites) have recently provided commercially viable and cost effective access to near space environments. Multiple launches of autonomous, spacecraft appear to be within reach, vastly expanding the future prospects of astrobionic research on small multicellular organisms. In this context, integrated LOC devices, due to their small small-size, can be prospectively designed to provide fully autonomous secondary payloads for studies of fundamental biological processes during space flight conditions. Current advances in MEMS can also support automated and miniaturized handling and analysis technologies along with life support capabilities and transfer of data to Earth. Such advancement and introduction of new functionalities will vastly expand the range of currently existing research opportunities for both terrestrial medicine and widely anticipated long-duration space missions. In light of apparent differences between conventional cytometric techniques, we postulate that a new term “wormometry” is needed to distinguish these up-and-coming technologies for the analysis of small model organism that go far beyond the conventional meaning of “cytometry.”

Acknowledgements

The authors thank Drs. David Strack and Julia Thompson (Union Biometrica, Holliston, MA, USA) for providing exemplary data on COPAS and BioSorter technologies; Dr. Pierre Delaage (Trophos SA, Marseille Cedex, France) for sharing materials on TROPHOS Plate RUNNER HD; Dr. Helmut Knapp (Microfluidics & Liquid Handling Section, CSEM SA, Alpnach, Switzerland) for generously providing data on proprietary CSEM ZebraFactor and XenoFactor technologies. The authors declare that views and opinions were not influenced by any conflicting financial interests.