Zebrafish embryo development in a microfluidic flow-through system

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Zebrafish embryo development in a microfluidic flow-through system†

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Eric M. Wielhouwer,‡a Shaukat Ali,‡a Abdulrahman Al-Afandi,a Marko T. Blom,b Marinus B. Olde Riekerink,b Christian Poelma,c Jerry Westerweel,c Johannes Oonk,b Elwin X. Vrouwe,b Wilfred Buesink,b Harald G. J. vanMil,a Jonathan Chicken,d Ronny van ’t Oeverb and Michael K. Richardson*a Received 24th September 2010, Accepted 17th March 2011 DOI: 10.1039/c0lc00443j The zebrafish embryo is a small, cheap, whole-animal model which may replace rodents in some areas of research. Unfortunately, zebrafish embryos are commonly cultured in microtitre plates using cellculture protocols with static buffer replacement. Such protocols are highly invasive, consume large quantities of reagents and do not readily permit high-quality imaging. Zebrafish and rodent embryos have previously been cultured in static microfluidic drops, and zebrafish embryos have also been raised in a prototype polydimethylsiloxane setup in a Petri dish. Other than this, no animal embryo has ever been shown to undergo embryonic development in a microfluidic flow-through system. We have developed and prototyped a specialized lab-on-a-chip made from bonded layers of borosilicate glass. We find that zebrafish embryos can develop in the chip for 5 days, with continuous buffer flow at pressures of 0.005–0.04 MPa. Phenotypic effects were seen, but these were scored subjectively as ‘minor’. Survival rates of 100% could be reached with buffer flows of 2 mL per well per min. Highquality imaging was possible. An acute ethanol exposure test in the chip replicated the same assay performed in microtitre plates. More than 100 embryos could be cultured in an area, excluding infrastructure, smaller than a credit card. We discuss how biochip technology, coupled with zebrafish larvae, could allow biological research to be conducted in massive, parallel experiments, at high speed and low cost.

Introduction In biological and biomedical research, there is an unmet need for low-cost, high-throughput in vitro animal models.1,2 Whole animal models are valuable because they provide data that may be extrapolated to humans; they also allow complex organismal functions (e.g. behavior and development) to be studied.3 On the other hand, in vitro models (e.g. cell and tissue culture) offer the advantages of low cost, of being less prone to legal and ethical restrictions and of having the ability to be scaled-up to highthroughput (1000–10 000 assays per day4) or ultra high throughput (100 000 assays per day5). Zebrafish (Danio rerio) embryos have been proposed as an in vitro animal model which could bridge the gap between simple assays based on cell culture, and biological validation in whole

a Institute of Biology, Leiden University, Sylvius Laboratory, Sylviusweg 72, 2333, BE, Leiden, The Netherlands. E-mail: richardsonmk@biology. leidenuniv.nl b Micronit Microfluidics BV, Enschede, The Netherlands c Laboratory for Aero & Hydrodynamics, Delft University of Technology, The Netherlands d FLIR Systems LTD, Nottingham, UK † Electronic supplementary information (ESI) available. See DOI: 10.1039/c0lc00443j ‡ These authors contributed equally.

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animals such as rodents.1 The zebrafish embryo has a small size, low-cost, rapid development and can be raised easily in large numbers. It also has a transparent body, which makes it relatively easy to collect numerous data points using high-quality imaging (including the fluorescence imaging of transgenic lines6). This species is therefore suitable not only for high throughput screens, but also for high content screens. Indeed it is already beginning to be used in large-scale screens and assays.6–8 Several zebrafishembryo assays can help to predict drug safety in humans,9,10 and zebrafish disease models have been developed.11,12 Unfortunately, the ambitions that biologists have for zebrafish embryos have outstripped the available culture protocols, borrowed as they are from traditional cell culture. Thus, zebrafish embryos are commonly raised for assay purposes in plastic microtitre plates or Petri dishes.7,13 In Table 1 we give examples of assay protocols in zebrafish studies. Typically, the buffer is refreshed periodically (‘static renewal’) or not at all (‘static nonrenewal’).14 Periodic aspiration and replacement of the buffer is extremely invasive, causing stress to zebrafish embryos and requiring enormous care in order to avoid embryos being damaged or sucked up. Another issue is that static replacement regimes may not be ideal for the zebrafish, a species which normally breeds in slow-flowing waters.15 The imaging of embryos in a microtitre plate is distorted, not only by the depth of the buffer filling the well, but by the curved meniscus of the buffer which Lab Chip, 2011, 11, 1815–1824 | 1815

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Table 1 A selection of assays to give an indication of protocols currently used in zebrafish research. In this case we list ethanol teratogenicity assays to show the wide variation in setup used for testing a single reagent. Note: dpf ¼ days post fertilisation; hpf ¼ hours post fertilisation Duration of exposure

Stage of exposure

Plate format

Ref.

Acute (1 h) Acute (2 h) Acute (3 h) Acute (1 h) Acute (1 h) Acute (20 min) Chronic Chronic (2 weeks) Chronic Chronic Chronic (3 d) Chronic (3 d) and acute (4 h) Chronic (6 h) Chronic (ca. 20 h) Chronic (6 d) Chronic (ca. 20 h) Chronic (ca. 20 h) Chronic (ca. 24 h)

3–4 month 1 dpf 256 cells, high, dome/30% epiboly, germ-ring 4 month 6 dpf 7 dpf 6–24, 12–24, 24–36, 48–60, 60–72 hpf Young adult 6–24, 12–24, 24–36, 48–60, 60–72 hpf 1 dpf 1 dpf 2 dpf 1 dpf 1 dpf 1 dpf 1 dpf 1 dpf 1 dpf

Aquarium (15 l) Petri dish, 60 per dish, tank, 20 per tank Petri dishes or glass beakers Tank 96-Well plate 10 per chamber 8  6  2 cm Petri dish 5-Gal aquarium Petri dish Aquarium 6-Well plate, 10 per well 6-Well plate Petri dish Petri dish 24-Well plate, 10 per well 5 ml (format not specified) Petri dishes or glass beakers Glass beaker

33 34 27 35 30 25 36 37 38 39 40 41 42 43 44 45 46 47

interferes with phase contrast and bright-field microscopy. A final problem with traditional, static replacement regimes is the possible bolus effect resulting from the intermittent replacement of buffer and its contained drug. An example of static non-renewal culture of zebrafish embryos is their successful growth inside Teflon tubing, each embryo being isolated in a drop of buffer.16 Chronic exposure to drugs is possible in such a system, but the embryo is not accessible during the experiment. Furthermore, culture in Teflon tubing involves distortion of the image because of the curved surfaces, and does not provide continuous buffer refreshment. Another study in which the microfluidic culture of embryos is described (mouse, in that case) used static culture in droplets with no fluidic flow.17 Another approach was developed by a student team and reported in an educational-themed issue of the journal Zebrafish. Unfortunately, no relevant biological data were given in that paper, although the authors claim that the zebrafish could survive for a few days in their single-well PDMS (polydimethylsiloxane) open set-up in a Petri dish.18 While this is a very intriguing report, the ‘biorector’ described does not appear to meet the criteria of a high throughput, microfluidic technology. We believe that true microfluidic technology could provide a quantum leap forward in this field by providing non-invasive culture conditions and high-quality imaging, as well as the ability to access the embryo at any stage of development (this not being possible, for example, with the culture of embryos inside lengths of tubing). For the purposes of our study, true microfluidic technology involves: continuous flow-through (‘dynamic renewal’) of pressurised buffer; the embryos being continuously accessible and isolated in parallel arrays to prevent crosscontamination; the use of small culture volumes to save valuable compounds; and the capability of high quality, real-time imaging of the embryo. Microfluidics have been used for sorting nematode (Caenorhabditis elegans) embryos into 96-well plates, and for holding nematode larvae.19 Unfortunately, no animal embryo has been shown to be capable of developing in a true microfluidic 1816 | Lab Chip, 2011, 11, 1815–1824

environment. Our aim here is to determine whether zebrafish embryos can indeed complete normal organogenesis under such conditions.

Results and discussion Properties of the biochip The microfluidic chip that we have designed and tested here (Fig. 1a) is made of three layers of bonded borosilicate glass, with an array of wells connected in parallel by channels. The temperature of the wells can be controlled by water flowing through in-built heating channels (Fig. 1b and c). Thermal imaging showed that the temperature difference between the extreme ends of any row (Fig. 1b and c) ranged from 0.08– 0.53
C at a flow rate of heating water of 1 ml per min per chip. Several prototypes were tested, having wells of either 1.5 mm, 1.67 mm, 1.83 mm or 2 mm inner diameter; the four well sizes were in either round or square shape, giving eight biochip prototypes in total. For comparison, we determined that the diameter of a pre-hatching zebrafish embryo (2–4 somite stage) averages 806 mm (SD, 31) or 1237 mm (SD, 24) including the chorion (N ¼ 15). Therefore the embryo fits comfortably into the well, and can swim around after hatching (Movies S1 and S2†), but is confined within a much smaller physical area than is the case with 96-well plates. Fluidic flow We next looked at how buffer circulated in a well containing an embryo. Obstruction of the buffer flow by the embryo, or by debris (including the chorion shed at hatching), is prevented by having 3 inlet and 3 outlet channels per well (Fig. 2a). Crosscontamination between wells (e.g. by chemicals or infectious agents) is ameliorated by the parallel arrangement of wells (Fig. 1a). The accumulation of air bubbles under the lid is prevented by having the outlet channels positioned high on the wall (Fig. 2b and c). In Movie S1†, several air bubbles can be seen to spontaneously shrink and disappear during operation of the chip. This journal is ª The Royal Society of Chemistry 2011

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Fig. 1 Layout and thermal properties of the biochip. (a) An example of a 32-well chip. The pre-heated water circulating in the temperature channels (tc) is completely isolated from the buffer in which the embryos develop. Buffer enters the chip via inlet (bi) and outlet (bo) ports, and circulates through a parallel array of feeder channels. These in turn are connected to the wells (w) by smaller channels, each of which is connected to the well by three terminal branches, so each well has a total of 6 connections (3 inflow, 3 outflow). Scale bar ¼ 1 cm. (b) Thermographic recording made with a FLIR SC5600-M large format infrared camera of a 24-well chip running in a room with 20
C ambient temperature. Heating water was circulated via thermal inlet (ti) channels and the common thermal outlet (to) channel. The temperature of heating water entering at ti1 was 26.06
C; ti 2, 26.20
C; ti 3, 25.04
C; ti4, 25.11
C. (c) Temperature profile from (b), showing that a stable temperature can be maintained in different wells of the chip. There difference along each row (i.e. between wells 1 and 6) averages 0.18
C. A stable temperature can be maintained in the chip even though the surrounding air is at room temperature.

We tested fluidic properties of the wells using micro-PIV techniques.20,21 The flow regime over the range of flow rates of 1.0–10.0 mL per well per min, was laminar, with no vortices or turbulent motion (Fig. 2d–i). This is predictable, given that the dimensionless Reynold’s number (Re ¼ rVD/h) is significantly smaller than unity, indicating that viscous forces dominate over any inertial effects. Interestingly, it can be seen in Movie S3† that twitching movements of the embryo caused periodic rearrangement of the flow pattern of buffer in the well. Embryo survival in the biochip We next wanted to determine whether zebrafish embryos could survive and develop in a microfluidic environment because this This journal is ª The Royal Society of Chemistry 2011

has never been shown for any animal embryo. As controls, we cultured zebrafish embryos in conventional 96-well microtitre plates with periodic (24 h) static renewal of buffer. We dispensed 8 hpf zebrafish embryos into the biochip, 1 per well. We plated them into the chip at 8 h, because this avoids the mortality-wave seen in earlier stages.22 We first looked at survival of embryos after 5 days of culture. We chose 5 days as the cut-off point in order to conform to local ethical requirements. However, at five days, most of the organs are developed and the larva already shows a complex behavioural repertoire. In the 96-well plate controls, there was 100% survival at 5 d. In the biochip, 5 d survival of the zebrafish was strongly influenced by the buffer flow-rate (Fig. 3). The average 5 d survival from all experiments at each flow rate was as follows: 36.7% survival at 0.5 mL per well per min; 53.9% at 1.0; 88.3% at 2.0; 87.5% at 4.0; and 71.1% at 6.0 mL per well per min. Survival in the biochip at 2.0–4.0 mL per well per min was significantly higher (p < 0.01 using one-way ANOVA) than at 0.5 mL per well per min. Considering single runs with individual chips, survival ranged from 19.0% (one chip run at 0.5 mL per well per min) to 100% (two chips run at 2 mL per well per min). In the biochip, well size and shape had no effect on survival (biochip wells of 1.67 mm or 1.83 mm inner diameters, round or square, were examined; twoway ANOVA). Interestingly, the hatching of embryos in the biochip (Fig. 3b) was suppressed at higher flow rates. To see whether oxygen shortage could explain the poor survival at 0.5 mL per well per min, we forcibly aerated the buffer reservoir using an aquarium air-stone. The average 5 d survival at 0.5 mL per well per min with no extra aeration was 36.7%; at the same flow rate, with forced aeration of the buffer, survival only slightly increased to 43.8%. Thus, forced aeration was not able to raise the survival rates to those seen at 2.0 or 4.0 mL per well per min, implying that some deleterious effect other than oxygen depletion is at work when low flow rates are used. Phenotypic screening of embryos Surviving embryos at 5 d were fixed, stained and screened morphologically (Fig. 3c and 4). The phenotypes scored are listed in Table 2, and the data given in Table 3. Embryos grown in the biochip had a shorter average body length (3.63 mm) compared to controls in 96-well plates (3.84 mm) (see Fig. 5a). This difference was highly significant (p > 0.001 using an ANCOVA linear regression model). Greater embryo-length was obtained for flow rates of 1, 4 and 6 mL per well per min respectively compared to 0.5 mL per well per min, the difference being very significant (p < 0.05, df 467; Box–Cox analysis followed by a factorial ANOVA performed on the squared embryo-length against buffer flow as an explanatory factor). To look for signs of stress, we examined the phenotype of melanin-containing pigment cells (melanocytes) at 5 days. Pigment dispersion is known to occur in teleosts exposed to stressors.23–25 We performed a cubic transformation of the ratio between the number of embryos with contracted pigment pattern, and the total number of embryos, and a two-way ANOVA for factors FLOW RATE and WELL-TYPE. No effect was found for FLOW RATE, but a significant effect was observed for WELL-TYPE, the ratio being 0.96 for 96-well Lab Chip, 2011, 11, 1815–1824 | 1817

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Fig. 2 Fluidic flow in the biochip. (a) Zebrafish embryo in a 2.0 mm (inner diameter) well to show the positioning and relative size of an embryo in the well (photographed without a lid, and therefore without buffer flow, so as to give an unobstructed view of the connections); tc, temperature channel; bi, buffer inlet, bo, buffer outlet. The central, darker ring (r) in the picture is the sandblasted wall of the well. (b) Schematic drawing showing the levels of the z-planes and x  y coordinates, and their relation to the inlet channels (blue) and outlet channels (orange). (c) Schematic diagram of flow pattern around and over the embryo. (d–i) micro-PIV recordings of flow patterns in a fluidic flow experiment using micro-PIV techniques.20,21 Polystyrene particles (1.3 mm diameter, containing rhodamine and coated with PEG) were used. The flow rate used in this experiment was 1 mL per well per min and the inner diameter of the well was 1.5 mm. (d, e, and f) schematic interpretations of the actual recordings (g, h, and i), respectively. Red arrows in g, h, and i indicate the most rapid flow, blue arrows the slowest, and blue dots represent flow parallel to the z-axis. bi, position of buffer inlet; bo, position of buffer outlet; r, wall of well.

plates versus 0.64 for biochips (p < 0.01, df ¼ 8). In summary, there is a significantly higher incidence of a putative stress phenotype (dispersed pigment granules) in embryos grown in the biochip. Other phenotypic effects were screened for, and we found mild yolk sac oedema (compare Fig. 4b and c) much more commonly in the biochip embryos than in controls (p < 0.001, generalized linear model). However, there was no significant difference in the incidence between biochip and 96-well culture (p > 0.05) of other malformations (cardiac oedema, pectoral fin hypoplasia, branchial arch hypoplasia, hypoplasia of Meckel’s cartilage, bent tail, and bent primary axis).

phenotypic effect in the zebrafish embryo;12,26 it passes easily through the chorion;27 and it is widely used as a carrier solvent28 in biomedical research. We scored the embryos at 5 d for a variety of phenotypic abnormalities (Table 2). Then, we classified each embryo as to whether it was mild, moderate or severe in terms of its clustering of phenotypic abnormalities, using the criteria in Table 4. In control (vehicle only) embryos there was a very low incidence of severely abnormal embryos (Table 5). But with ethanol exposure, the percentage of severely abnormal embryos increased in both the 96 well plate (to 85%) and the biochip (to 65%). Implications of the current findings

Acute ethanol exposure test To see whether we could introduce bioactive compounds into the wells during an experiment, we introduced 10% ethanol into the inflow stream of the biochip for 1 h at prim-16 stage. Ethanol was chosen because of several reasons: it produces a strong 1818 | Lab Chip, 2011, 11, 1815–1824

It is not clear what causes the minor phenotypic effects (e.g. yolk sac oedema, slight reduction in body length) in biochip-raised embryos. Possible explanations could be physical features of the biochip itself, such as the small size of the wells. Fig. 4a and Movie S2† show that there is sufficient room for the embryos to This journal is ª The Royal Society of Chemistry 2011

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Fig. 3 Data on survival and hatching percentage of embryos cultured in the biochip. (a) 5 d survival in the biochip, plotted as a function of buffer flow rate. The results from 4 different chip designs (each with wells of a particular size and shape) are shown. For each data point, N ¼ 32 (embryos, except those for the biochip experiments with a flow rate of 1 mL per well per min, where N ¼ 64). For each experiment, a control was run simultaneously in a 96-well plate, placed next to the biochip and using embryos of the same stage and from the same mating. For each control, N ¼ 32 embryos. (b) Hatching rate as a percentage of surviving embryos at different time points and at different flow rates of buffer (in mL per min per well). Higher flow rates in the biochip tend to delay or suppress hatching, relative to controls. Each error bar represents  SEM (standard error of the mean) of N ¼ 48 embryos, that is, 4 replicate chips, with 12 embryos each, per flow rate; for the 96-well plate controls, N ¼ 160 embryos, consisting of 5 replicates each with 32 embryos. (c) Incidence of malformations in surviving embryos at 5 days in 96 well plates and biochips (each line representing a different flow rate). The number of embryos affected by various malformations is shown (note that some embryos had multiple malformations and therefore the numbers do not add up to 100%). Note also that the 96-well plates had daily static replacement regimes and so the different ‘flow-rates’ indicated for 96-well plates are simply referring to the biochip run for which they were the control. Key: pf, pectoral fin hypoplasia; bt, bent tail; ys, yolk sac oedema; pc, pericardial oedema; Mc, Meckel’s cartilage hypoplasia; normal, none of the other malformations in this series were seen; ba, branchial arch hypoplasia; bc, curvature of body axis.

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Fig. 4 Embryos cultured in the biochip. (a) Consecutive photos of the same embryo developing in the same well (1.8 mm internal diameter) of a 32 well biochip with a buffer flow of 2 mL per min per well (note that by 4 days, the embryo had swum into a different focal plane in the upper part of the well). Each picture is framed by a circular hole in the metal clamp that holds the lid in place. Notice that between 1 dpf and 2 dpf, the embryo has changed position within the chorion. (b and c) Two embryos grown in the biochip and fixed at 5 days (Alcian blue staining and clearing); ys ¼ yolk sac; (b) morphologically normal embryo; (c) embryo with mild yolk sac oedema. Scale bar in a ¼ 1 mm; scale bar in b and c ¼ 500 mm.

lie straight and swim around until at least 4 days of culture. Nonetheless, the embryo has approached the limits of the well at 5 d and this is perhaps an explanation for the increase in the putative stress phenotype (dispersed melanocytes) seen in the biochip embryos at that time. The ethanol exposure experiment shows that compounds can be introduced into the buffer stream at will and can produce an effect on embryo development. The percentage incidence of severe abnormalities after ethanol exposure in the biochip was less than that after the same test conducted in a 96-well plate. A possible explanation is that the specific gravity of ethanol is lower than that of water, resulting in a failure of the ethanol to enter the lower part of the well where the embryo is located. What are the potential advantages and limitations of microfluidic embryo culture? Our study suggests several advantages for Table 2 Phenotype analysis. Description of the seven categories used to score larval phenotype at 5 dpf Larval phenotype

Criteria

1. Normal

Absence of any of the phenotypes listed below: Dispersed melanocytes Presence of pericardial oedema Presence of yolk sac (vitelline) oedema Meckel’s cartilage grossly hypoplastic, missing or unfused in midline. These effects may be unilateral or bilateral One or more cartilages of the branchial skeleton hypoplastic or missing One or both pectoral fins hypoplastic or missing

2. Pigmentation 3. Heart 4. Yolk 5.Meckel’s cartilage 6.Branchial arches 7. Pectoral fins

1820 | Lab Chip, 2011, 11, 1815–1824

medical and life sciences research. In the field of highthroughput, high content research (i.e. when very large numbers of samples are screened, and a large number of data points is collected) a biochip could represent a major advance over conventional microtitre plates. First, the biochip concentrates embryos into a small physical area, because the wells are both Table 3 The effects of flow rate in the biochip on survival and phenotypes of embryos at 5 dpf. For each flow rate, a 96-well plate control with static replacement of buffer was established. Biochip data for each flow rate are pooled from chip versions with differences in well diameter. This is because the statistical analysis revealed no significant effect of well size or shape on survival (see main text) Survivors Flow rate/ Total Dead Losta (5 dpf)b Normal Abnormalc mL per min Well N (%) N (%) N (%) N (%) N (%) per well format N 0.5 1.0 2.0 4.0 6.0

96-well Chips 96-well Chips 96-well Chips 96-well Chips 96-well Chips

32 128 32 128 32 128 32 128 32 128

0 (0) 81 (63) 0 (0) 92 (72) 0 (0) 15 (12) 0 (0) 16 (13) 0 (0) 37 (29)

0 (0) 5 (4) 0 (0) 3 (2) 1 (3) 12 (9) 2 (6) 8 (6) 3 (9) 10 (8)

32 (100) 42 (33) 32 (100) 33 (26) 31 (97) 101 (79) 30 (94) 104 (81) 29 (91) 81 (63)

29 15 31 11 24 37 23 17 20 20

(91) (36) (97) (33) (77) (37) (77) (16) (69) (25)

3 27 1 22 7 64 7 87 9 61

(9) (64) (3) (67) (23) (63) (23) (84) (31) (75)

a ‘Lost’ indicates that embryos were lost during processing (mostly through aspiration during pipetting of buffer or other reagents). b Phenotype at 5 dpf was classified as normal or abnormal according to the criteria in Table 2. c Abnormal embryos were classified as mild, moderate or severe according to the criteria listed in Table 4.

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Fig. 5 Further characterisation of the phenotypes of embryos grown in the biochip versus controls grown in 96-well plates. (a) Chart of body length at 5 dpf for surviving embryos grown in the biochip (pooled data from all biochip version 1 models) versus their respective 96-well plate controls (which had static volumes with no buffer flow). The abscissa gives different flow rates per well in the biochip. The number inside the base of the bars ¼ N embryos. (b) Relative incidence among 5 dpf survivors of dispersed versus contracted melanocyte morphology.

small and close together. This facilitates parallel imaging of multiple wells at high resolution. It also reduces the ‘seek’ time needed to locate embryos, either manually or automatically. Furthermore, there is no water meniscus in the biochip to distort the image, and the walls are frosted, avoiding mirroring artefacts. Finally, the biochip is made of optical-quality glass, and the depth of fluid covering the embryos is much less than that in a 96-well plate. As shown in Movie S4† it is possible to obtain high quality imaging of zebrafish lines in the chip. With respect to buffer replacement, the great advantage of a microfluidic flow-through system is that there is no repeated invasion and disruption of the embryos environment by draining This journal is ª The Royal Society of Chemistry 2011

and refilling. This is crucial because zebrafish embryos are sensitive to handling stress.29 The small volume of the biochip wells (ca. 10 ml) means that some types of experiment, particularly acute drug exposures, will consume far less precious reagent or drug than the same experiment in microtitre plates (for example, 250 ml is typically used per well). This makes the biochip promising for drug discovery where only small quantities of compound are available, or where the compound is very expensive. The most obvious limitation of microfluidic systems is size; at some point, the embryos will simply outgrow the wells. This is not, of course, a problem in short-term assays up to 4 or 5 d. It Lab Chip, 2011, 11, 1815–1824 | 1821

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Table 4 Severity scale used to express the degree to which individual embryos were phenotypically abnormal. Branchial and Meckel’s cartilage defects were excluded from the ‘moderate’ category because craniofacial defects are typical of severe ethanol teratogenicity in the clinical situation.48 Severity

Criteria

Mild

An individual embryo had a single abnormality (of any type listed in Table 2) An individual embryo had any two defects, excluding branchial and Meckel’s cartilage abnormalities (i.e. the embryo showed two from categories 2–4, or 7, in Table 2) An individual embryo had alteration of the branchial arches and/or Meckel’s cartilage combined with one or more of other defects (2–7 in Table 2)

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Moderate

Severe

Experimental Biochip design The biochip prototypes were specially fabricated for this study by Micronit Microfluidics (Enschede, The Netherlands). They consists of three layers of bonded borosilicate glass into which an array of channels and wells is introduced by etching and powderblasting. A range of prototypes was tested, with slight variations in design (round or square cross-sections, and well sizes of 1.5– 2.0 mm inner diameter). Temperature channels permit the circulation of pre-warmed water through the chip. The open wells can be closed by a sandwich of a silicon polymer sheet, and a layer of glass, the whole assembly being compressed in a holder to make it watertight.

Embryo preparation should also be noted that the small well size of the biochip does not necessarily mean a saving of test compound—if chronic exposure is used for the full 5 days of embryo culture. For example, with a buffer flow rate of 2 mL per well per min, the biochip consumes 14.4 ml of buffer per embryo over 5 days. This compares with 2.35 ml per embryo in a 96-well plate for the same time period, assuming daily buffer replacement (see Methods for buffer refreshment protocol). In such cases, conventional 96-well zebrafish assays, which have now a high level of automation,13 remain a valuable alternative. In conclusion, we show that zebrafish embryos can develop normally in complete isolation with constant buffer flow and strictly controlled environmental parameters. Crucially, the zebrafish embryos can undergo normal organogenesis in a pressurized fluid stream. This is important because pressurized flow distinguishes microfluidics systems from conventional culture systems. Problems that need to be solved include the rather high occurrence of mild yolk sac oedema in the biochip embryos. Ultimately, it is possible that the microfluidic culture of vertebrate embryos could become a bridge between conventional cell culture, and whole animal models. Such a bridge is especially important, given the need to find alternatives to mammalian whole-animal experiments; and the need to find new medicines by means of more efficient assays.

Adult zebrafish (Danio rerio) were maintained in aquaria at 26
C under a cycle of 14 h light, 10 h dark. Eggs were obtained by random pair-wise mating. The eggs were transferred to 9 cm Petri dishes containing 0.1 Hanks’ Balanced Salt Solution30 (0.1 HBSS) at pH 7.46, and periodically rinsed to remove debris and dead embryos. The HBSS did not contain methylene blue or antibiotics.

Experimental setup All biochip culture experiments were carried out at 28.0  0.5
C under a 14 h/10 h light/dark cycle. Embryos of 8 h postfertilisation (hpf) were loaded, one per well, with intact chorion, into the biochip with residual buffer. Then, the lid was sealed (see above), the flow of buffer (0.1 HBSS) was initiated, and the setup ran continuously until the embryos had reached 5 dpf. Inlet and outlet channels of the wells were fed with the buffer from a high-performance liquid chromatography (HPLC) pump, without a de-gasser. The biochips were connected to the pump in parallel by means of phenyl/methyl deactivated capillary tubing (150 mm inner diameter and 375 mm outer diameter; BGB Analytik AG: Schlossboeckelheim), and cross interconnectors.31 The buffer reservoir was not actively aerated but had a loose-fitting foil cover. Buffer was not recirculated.

Table 5 General outcomes of ethanol treatment. Overview of total number embryos treated, survival at 5 dpf, the presence of morphological abnormalities at 5 dpf, and the degree of severity of those abnormalities Morphology (5 dpf)a Severity of abnormalityb Well format

Treatment

Total N

96-well

Veh EtOH Veh EtOH

96 96 64 64

Chip (1.5 mm, square)

Dead N (%)

Lostc N (%)

Survivors (5 dpf) N (%)

Normal N (%)

Abnormal N (%)

Mild N (%)

Moderate N (%)

Severe N (%)

2 (2) 70 (73) 13 (20) 23 (36)

26 (27) 6 (6) 12 (19) 4 (6)

68 (71) 20 (21) 39 (61) 37 (58)

35 (51) 0 (0) 11 (28) 5 (14)

33 (49) 20 (100) 28 (72) 32 (86)

20 (61) 3 (15) 9 (32) 1 (3)

11 (33) 0 (0) 15 (54) 7 (22)

2 (6) 17 (85) 4 (14) 24 (75)

a Morphology at 5 dpf was classified as normal or abnormal according to the criteria in Table 2. b Abnormal embryos were classified as mild, moderate or severe according to the criteria listed in Table 4. c ‘Lost’ indicates that embryos were lost during processing (mostly through aspiration during pipetting of buffer or other reagents).

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Controls

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To compare performance of the biochip with conventional culture conditions we made control cultures using zebrafish embryos cultured in conventional 96-well microtitre plates. Embryos of 8 hpf, randomly selected from the same batches used for the biochip experiments, were established in 96-well microtitre plates (Costar 3599, Corning Inc., NY). One embryo was placed in each well, with 250 mL 0.1 HBSS. The buffer was replenished every 24 h. The daily refreshment of buffer was done by replacing 175 mL, three times. The buffer, temperature and light cycle were the same as described above for the biochip experiments.

biochip commonly showed minor phenotypic effects—including the possible stress phenotype of dispersed melanocytes. However, there was no increase in gross malformations. Our scoring of the former as ‘minor’ defects is of course subjective, and does not rule out the possibility of significant but undetected effects. There was a strong, non-linear relationship between buffer flow-rate and 5 d embryo survival in the biochip, such that the optimal flow rate was in the range 2.0–4.0 mL per well per min. Survival rates at 5 d reached 100% in two chip runs at 2.0 mL per well per min. These results could lead to a new generation of assays for the pharmaceutical industry based on the low-cost, microfluidic culture of zebrafish embryos.

Acute ethanol exposure Embryos were established in the biochip (1.5 mm internal well diameter, 1.0 mL per well per min flow rate), and in 96-well plates (controls) as described above, except that the embryos used were all at the prim-16 stage32 (approximately 1.5 dpf). This is because preliminary data indicated that this was an ethanol-sensitive stage. Embryos were exposed for 1 h to 10% ethanol (1.64 M in 0.1 HBSS) or buffer only as controls. The ethanol was high purity, medical grade (Emprove ethanol, Cat. No. 100971, Merck KGaA, Darmstadt, Germany). Exposure to ethanol in the biochip was accomplished by temporarily disconnecting the normal buffer reservoir, and connecting in its place, for 1 h, the 10% ethanol/buffer reservoir. Embryos were not dechorionated since the chorion is known to be completely permeable to ethanol.27 The ethanol exposure was followed by rinsing with fresh 0.1 HBSS (4 rinses in the case of 96-well cultures). All embryos were then further cultured as described above, until five days. Analysis of embryos and statistical analysis design Total body length was measured from the rostral margin of Meckel’s cartilage in Alcian blue stained embryos, to the caudal extremity of the caudal fin fold. Statistical analysis for Fig. 3c was made in R. The need for data transformations was assessed by post-diagnostic and Box–Cox analysis of the model. Count data were analysed as a generalized linear model whereas continuous data as ANCOVA model. Fixation and staining of embryos At 5 dpf, embryos were fixed overnight in 4% paraformaldehyde and stained with Alcian blue as follows: embryos were rinsed 5 times in distilled water and dehydrated in a graded series of ethanol (25%, 50%, and 70%) for 5 minutes each. They were then rinsed in acid alcohol (1% concentrated hydrochloric acid in 70% ethanol) for 10 minutes and placed in filtered Alcian blue solution (0.03% Alcian blue in acid alcohol) overnight. Embryos were then differentiated in acid alcohol for 1 h, washed 2  30 min in distilled water. Finally, they were cleared and stored in 100% glycerol.

Conclusions We show for the first time that an animal embryo can develop in a microfluidic environment. Zebrafish embryos grown in the This journal is ª The Royal Society of Chemistry 2011

Acknowledgements We thank Jan de Sonneville and Dr Maxim E. Kuil for helpful discussions on imaging, microfluidics applications and for comments on the manuscript; Ewie de Kuyper, Arjen C. Geluk, Jeroen Mesman, Frits (M.W.) van Tol, and Tim A. P. Sanders helped design and build prototype biochip holders; Peter Steenbergen and Ulrike Nehrdich for culturing zebrafish embryos; Edwin Heida for scientific illustration; Nathan D. Lawson and Brant M. Weinstein for transgenic Fli1 eGFP zebrafish; Michael Richardson gratefully acknowledges the financial support of the Smart Mix Program of the Netherlands Ministry of Economic Affairs and the Netherlands Ministry of Education, Culture and Science.

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