Chip Design and Operation
The 3D chip was fabricated in a biologically compatible and optically transparent PMMA polymer using infrared laser micromachining. The chip encompassed seven separate layers assembled in two distinctive units: (i) the main structure formed by five top layers and (ii) heating module housing composed of two bottom layers. Layers in both units were optically aligned and thermally bonded (Fig. 1). Main units were then reversibly assembled using M2 stainless steel bolts. The chip design consisted of seven integrated modules: (i) a main channel (1.7 mm × 1.5 mm × 55 mm) for embryo loading and recovery, (ii) a linear array of 16 miniaturized traps for single embryo trapping and immobilization, (iii) a suction manifold (0.7 mm in depth) that created a drag force to immobilize embryos, (iv) a direct drug delivery channel (1 mm × 0.5 mm × 120 mm) that provided independent drug delivery to all embryo traps, (v) pumping unit with housings for two on-chip piezoelectric micropumps, (vi) thermistor probe housing located next to embryo trapping array and separated by a thin wall (0.2 mm), and (vii) a heating manifold housing for the ITO heater (Fig. 1). The design of the latter module allowed for temperature stabilization around the embryo-trapping array and provided heating of the drug solution flowing in the drug delivery manifold. The air pocket and thin PMMA wall (1 mm) between the heating manifold and the main chip unit facilitated the rapid heat transfer directly to immobilized embryos and drug solution.
To achieve trapping, culture and analysis of zebrafish embryos, a linear array of traps was ablated using laser raster mode to the depth of 1.6 mm (Fig. 1). Each trap had a conical geometry with a diameter of 1.8 mm on the top plane and a diameter of 1.5 mm on the bottom plane. Subsequently, channels with a diameter of 0.5 mm were laser drilled on the bottom plane of each well using a 50 μm infrared laser cutting beam. These channels are seen as small apertures at the bottom of microwells and interconnect the trapping array with a suction manifold located underneath (Fig. 1). The drug delivery manifold was ablated using laser raster mode to the depth of 0.5 mm, and its outputs allowed for simultaneous delivery of drugs to every trapping well (Fig. 1). The drug delivery manifold could be actuated independently from both the main channel and suction manifold.
The chip-based device featured two embedded piezoelectric ultrasonic microdiaphragm pumps that realized adjustable and linear flow rate between 0.01 and 3 ml/min (Figs. 1 and 2). Pumps were independently controlled by external microcontroller. In this regard, the chip was operated in two distinctive pumping regimens: (i) embryo loading and immobilization—No. 1 piezopump ON and connected to suction manifold outlet providing fluid negative pressure at the bottom plane of the trapping array to support embryo loading and trapping; (ii) embryo drug perfusion—No. 1 and No. 2 piezopumps ON—No. 1 piezopump delivering solution of the drug whereas No. 2 piezopump providing fluid negative pressure at the bottom plane of the trapping array to support uniform drug microperfusion around immobilized embryos (Figs. 1 and 2).
Zebrafish Embryo Trapping and Culture on a Chip
The chip design allowed for both single embryo occupancy in the traps and unobstructed passage of other embryos in the rectangular main channel following docking. The trapping principles exploited the combined gravitational-induced sedimentation of embryos and a low-pressure suction at the bottom plane of the device to rapidly attract embryos into the traps (Fig. 3). The zebrafish embryos have a substantial mass of up to 1 mg, and in the presence of gravity the embryos were deflected under the combined effect of suction flow and gravity toward the aperture of the traps (Fig. 3). The embryos falling into traps were then immobilized by a continuous suction at the bottom plane of the device (Fig. 3).
Figure 3. Embryo trapping and docking principles. (A) System exploits combined gravitation-induced sedimentation and low-pressure suction at the bottom plane of the device. The red arrows indicate the direction of the gravitational force, whereas the blue arrows indicate the suction flows. (B) 3D cartoon depicting the process of embryo trapping. (C) 3D streamlines of flow through the trapping array colored by velocity (m/s) obtained by computational fluid dynamics simulations depict the principles of fluid flow during embryo immobilization. (D) Embryo trapping efficiency characterized by the number of embryos captured divided by the number of embryos injected into the LOC device. Experiments were performed at varying flow rates in the main channel and suction manifold as indicated. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Subsequent embryos introduced into the device rolled freely on top of the immobilized embryos toward the next available trap (Fig. 3). The process was repeated until all traps were occupied, and the loading was then discontinued. After computational fluid dynamics simulations that predicted trapping efficiency close to 100%, experiments were performed to validate these assumptions (Fig. 3). Trapping efficiency was then calculated as the number of captured embryos divided by the number of embryos injected into the LOC device.
During experimental validation, the system achieved 100% trapping efficiency when actuated at a total flow rate of up to 0.6 ml/min (Fig. 3). Deterioration of the trapping efficiency at higher flow rates was also observed (Fig. 3). This was attributed to increased embryo velocities and high momentum that could not be compensated by suction-assisted trapping (Fig. 3). Importantly, over 99% of trapped embryos retained their position during the course of 72 hrs experiments, with no dislodgement observed when drug delivery manifold was actuated.
After trapping and extended culture with control E3 medium, we observed normal and uniform development of all embryos immobilized across the array of all traps. The normalized cumulative survival of embryos perfused at a total flow rate ranging from 0.1 to 1 ml/min was over 95% ± 5% (Fig. 4). The embryo development and viability was uniform across the traps 1–16. Furthermore, developing embryos reached all developmental staging criteria that were statistically comparable with static Petri dish control experiments. Interestingly, the cumulative survival of embryos considerably deteriorated when chip was actuated at flow rates lower than 0.1 ml/min or when chip perfusion was disengaged (no-flow conditions) (Fig. 4). The decreased survival at very low flow rates was associated with a depletion of oxygen inside the chip when insufficient exchange of medium inside the gas nonpermeable PMMA device was present.
Figure 4. Development of zebrafish embryos in a chip-based device. (A) Brightfield image depicting Tg(fli1a:EGFP) embryos at 60 hpf. Note normal development with fully pronounced head, eyes, tail, and melanocytes. (B) Fluorescence image of Tg(fli1a:EGFP) embryos shown in (A) at 60 hpf. Note developing normal vasculature and central nervous system (green fluorescence). (C) Viability of WT and Tg(fli1a:EGFP) embryos as a function of flow rate. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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Proof-of-Concept Automation Interface
For the LOC field and miniaturized bioanalysis to become mainstream laboratory technology, key engineering challenges still need to be addressed. These include, among others, both on-chip and off-chip integration and simplification of many functional components, such as excitation and collection optics, automatically actuated reagent reservoirs, and miniaturization of fluidic actuators such as pumps and valves . So far limited progress has been made in user-friendly integration of LOC with the macro-world. Design of the innovative off-chip interfaces and automation of many functional mechanisms to provide truly turnkey and automated microfluidic devices is, however, of utmost importance for the realization of true micrototal analysis systems.
In this regard, we demonstrated a preliminary design of a new off-chip automation interface capable of automating most of the experiment procedures to achieve higher throughput, lower cost, and low turnaround time for the zebrafish embryo tests (Fig. 2). The platform is designed to load embryos into the chip-based device, control liquid perfusion, maintain the microenvironment permissive for embryo development within the device, and perform time-resolved fluorescent imaging of developing transgenic zebrafish embryos. Figure 2 represents an overview of the interface and its size comparison with conventional stereomicroscope.
The system was equipped with four main modules: (i) a robotic servo actuator-driven one-directional stage that holds integrated chip-based device; (ii) a micro-servo-driven pinch valve for rapid fluid control and drug switching; (iii) ARM-architecture microcontroller handling control over stage movements and pinch valve operations; and (iv) miniaturized USB fluorescent microscope with integrated array of blue LEDs (510 nm emission), excitation filter, and fully programmable time-resolved data acquisition (Fig. 5).
Figure 5. Comparison between resolution and sensitivity achieved by a conventional fluorescence stereomicroscope (Nikon SMZ1500 equipped with DS-U2/L2 camera) and a miniaturized fluorescent UB microscope (1.3 MP AM4113T-GFBW Dino-Lite Premier). ×4 magnification was digitally zoomed in for reference. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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In this work, we concentrated on evaluation of miniaturized and automated fluorescence image acquisition system that could substitute bulky and expensive conventional fluorescent microscopes (Fig. 5). The target application was capability to perform time-resolved imaging with every image collected every 60 min. Moreover, the CMOS sensor and dedicated optics should provide enough resolution and sensitivity to detect characteristic patterns of intersegment vessels within developing zebrafish embryo (Figs. 4 and 5). In this context, we have integrated into our system a USB-powered miniaturized fluorescence microscope (Dino-Lite). The proprietary software was fully capable of performing time-lapse imaging with automatic and integrated control over LEDs and exposure adjustments. We have found that the LED-based excitation coupled with built in collection optics and a 1.3-megapixel color noncooled CMOS sensor (SXGA) was adequate to collect fluorescence signals at both low and higher magnifications (Fig. 5). At low magnification, the resolution was comparable with that achieved by a Nikon SMZ1500 fluorescent stereomicroscope equipped with a DS-U2/L2 camera (Fig. 5). However, at higher magnification necessary for pattern analyses of developing vessel formation, the 1.3-megapixel color CMOS sensor was found insufficient for acquiring clear intersegment vessel (ISV) images despite its excellent fluorescence sensitivity (Fig. 5). The vessels were still observable, but analysis at even higher magnification would require an integration of a CMOS sensor with higher resolution.
As the resolution of the images acquired on different systems depends on combination of (i) optical resolution of the lenses used and (ii) the digital resolution of the sensors, it is interesting to note that both USB microscope and stereomicroscope featured objective lenses with similar numerical apertures, 0.1 and 0.135, respectively. We conclude that the major limiting factor of the USB microscope was indeed its digital resolution of 1.3 MP compared with 5 MP camera mounted on a conventional stereomicroscope. Furthermore, the 1.3 MP unit was a color and noncooled sensor whereas conventional microscope sensor was a black and white sensor dedicated to acquisition of faint fluorescent signals and therefore equipped with a Peltier cooling module. Accordingly, the work is currently ongoing to develop a similar miniaturized and portable microscope system equipped with a 5-megapixel color CMOS sensor. It will be interesting to evaluate whether increase in sensor pixel numbers without incorporation of any cooling module will provide improvements in resolution and sensitivity. If successful, this will enable proliferation of new imaging capabilities at a fraction of cost, power usage of conventional microscopes while also providing substantial space savings, and user-friendly operation.
The presented reduced functionality prototype brings the future LOC system for analysis of small model organism a step closer to realization of complete analytical automation. The system was made as a functional prototype based on the use of standard microprocessors, off-the-shelf components, and de-centralized control over several modules. We anticipate that next stage of prototyping will involve complete integration of individual system components and centralized field-programmable gate array-based extended implementation that will integrate all critical hardware and software components of the system in a single field-programmable gate array chip. The work is on-going on miniaturized and hardware encoding system that will synchronize automatic stage movements with the image acquisition process from every single embryo immobilized in the trapping array. Moreover, development of customized automatic image acquisition algorithms and real-time image analysis will improve the throughput providing instantaneous quantification of acquired results.
In Vivo Angiogenesis Assays in Microfluidic Environment
Zebrafish has recently been reported as a very convenient tool to perform accelerated in vivo screening of new pharmacologically active compounds [4-6]. For instance, Tg(fli1a:EGFP) line expressing enhanced green fluorescent protein in endothelial cells represents a rapid biotest to visualize development of characteristic patterns of ISVs [6, 14-16]. The biotest principles are based on diffusion of small drug molecules into the embryo and induction of dose-dependent inhibition of ISV formation. The latter can be microscopically visualized as reduction of fluorescence signal and quantified to provide dose response analysis. Moreover, the zebrafish biotest performed on intact and developing embryo can at the same time provide valuable data on potential adverse drug effects based on the presence of discernible phenotypic effects such as: tail detachment, tail and fin morphology, accumulation of melanocytes, eye and lens formation, kidney development, and cardiovascular function (heart rate and blood flow).
In this context, we performed preliminary analysis of the applicability of the microfluidic embryo array for the analysis of small-molecule compounds with antiangiogenic properties using the transgenic zebrafish line Tg(fli1a:EGFP). The embryos were loaded onto a chip-based system at 16 hpf stage before sprouting of ISVs had begun. The embryos were then continuously perfused in a closed-loop perfusion at a flow rate of 0.2 ml/min with E3 media containing VEGFR1-3 inhibitor AV951 (Tivozanib) or VEGFR2/PDGFRβ inhibitor Sunitinib (Fig. 6). Images of developing embryos immobilized on chip-based devices were acquired at 0, 24, and 48 hrs intervals that corresponded to 16, 40, and 64 hpf developmental stages (Fig. 6). The control embryos perfused with dimethyl sulfoxide vehicle control exhibited a normal pattern of complete ISV formation over 48 hrs of culture on a chip-based device (Fig. 6). VEGFR1-3 inhibitor Tivozanib proved to be the most potent drug achieving 100% of ISV growth inhibition at 1 μM concentration (Fig. 6). The VEGFR2/PDGFRβ inhibitor Sunitinib achieved no inhibition at 1 μM concentration and only a partial inhibition of ISV when used at 25 μM concentration (Fig. 6). Convenient arraying of embryos in predefined spatial locations substantially accelerated the data acquisition in contrast to conventional Petri dish assays. Moreover, the drug exchange and delivery were performed automatically within seconds without embryo dislodgement or need for repetitive pipetting. This was particularly beneficial when embryos were pulsed with a 0.2 mg/ml of tricaine mesylate to provide anesthesia and inhibit the intrinsic embryo movements before image acquisition. The data provide a proof-of-concept that microfluidic devices can be readily applied to perform accelerated in vivo analysis on developing transgenic zebrafish embryos.
Figure 6. On-chip angiogenesis assay. Fluorescence imaging of Tg(fli1a:EGFP) embryos at 64 hpf. Transgenic embryos were arrayed and immobilized at 16 hpf and continuously perfused with E3 media containing vehicle control (dimethyl sulfoxide) or selected small-molecule antiangiogenic drugs (Sunitinib and Tivozanib). Right panel: microscopic visualization of patterns of ISV. White arrows: normal ISV growth; blue arrows: partial ISV growth inhibition; and red arrows; complete ISV growth inhibition. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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On-Chip Heavy Metal FET Test
Zebrafish has been extensively used as a model organism in various studies that assess toxicity endpoints of pollutants . In recent years, zebrafish embryo tests gained popularity as surrogate tests to ethically controversial fish acute toxicity tests [18, 19]. Standard FET test is conducted under static conditions in multiwell microtiter plates, but this method is potentially inadequate because of adsorption, degradation, and accumulation of wastes that may severely restrict the exposure to the chemical of interest. Flow-through type devices are therefore crucial for the careful assessment of toxicity when zebrafish embryo is used as a target model .
Respectively, the microfluidic embryo array was therefore used to perform preliminary toxicity tests for the analysis of heavy metals dissolved in zebrafish medium. Static FET tests were carried out according to DIN standard on 24-well microtiter plates as control experiments to find the “lethal concentration 50” value of copper sulfate solution, which was found to be 25 µM. The zebrafish embryos were sorted to fertilized and unfertilized embryos, and fertilized embryos were loaded onto a chip at 6 hpf stage. The chip was then perfused in a closed-loop perfusion at the flow rate of 0.2 mL/min with 25 µM copper solution diluted in E3 embryo culture media (Fig. 7). Images of developing embryos immobilized on chip-based devices were captured at 0, 3, 6, and 24 hr intervals that corresponded to 6, 9, 12, and 30 hpf (Fig. 7). Static FET tests were always carried out in a 60-mm Petri dish at the same concentration for comparison with experiments run in triplicate to increase the sample number. The static FET tests with 25 µM copper sulfate have exhibited a 33.3% mortality rate. Interestingly, the mortality rate of on-chip cultured embryos showed a significant increase to average of 69% observed in static FET experiments (Fig. 7). We conclude that this increase could be attributed to a rapid exchange of copper surrounding the embryo by continuous perfusion in the microfluidic environment. This has led to a vast reduction in adsorption, degradation, and accumulation of wastes that may severely bias the conventional static ecotoxicology assays. Moreover, in contrast to the conventional FET assays, the automated arraying and long-term immobilization of embryos on chip-based device have substantially amplified the convenience of handling and image acquisition. These data provided evidence that the microfluidic devices can be readily applied to perform FET assay on developing zebrafish embryos.
Figure 7. On-chip fish embryo toxicity (FET) assay. (A) Brightfield image of WT embryos at 0 and 24 hr stage since the initiation of copper exposure. Red arrows indicate the copper solution flow to the embryos. Note the clear difference between the alive and dead embryos at 24 hr stage. (B) Embryo mortality rate in different FET conditions performed on a chip-based device and in static control conditions. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
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