Microfluidic trap array for massively parallel imaging of...

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol

nature protocols | VOL.8 NO.4 | 2013 | 721

IntroDuctIonThe fruit fly, Drosophila melanogaster, is a well-established model organism of developmental genetics. As the Drosophila genome is 44% homologous with humans, fruit fly development is a use-ful platform for investigating questions in tissue engineering, as well as in developmental and age-related diseases1,2. In particu-lar, the mechanism by which cellular identities are determined in the development of Drosophila embryos can provide insight into the conserved developmental processes and help identify suffi-cient conditions for differentiation and tissue generation in tissue engineering. Diffusible molecules that act in a concentration- dependent manner, so-called morphogens, form the basis through which embryonic patterning is established and gene networks are regulated3. Maternally deposited Bicoid and Dorsal, for example, are morphogens that pattern the anterior-posterior and dorsal-ventral axes of the developing embryo, respectively, and set off a cascade of gene regulatory events necessary for specifying the various tis-sues4–7. Because of the ellipsoidal shape of the Drosophila embryo, studying gene expression patterns along the longer anterior- posterior axis is inherently better suited for microscopy, whereas imaging the shorter dorsal-ventral axis patterning is more cumber-some. Conventional dorsal-ventral gene patterning studies require the individual placement of embryos in an ‘end-on’ orientation (i.e., with the dorsal-ventral axis parallel to the cover glass), which severely reduces the throughput of such investigations.

Recently, we developed a microfluidic device for the high-throughput orientation of D. melanogaster embryos to enable facile, large-scale investigation of dorsal-ventral gene expression pattern-ing8. Through the use of passive hydrodynamics, embryos can be oriented in an end-on position and subsequently immobilized in what we refer to as ‘embryo traps’, which are basically embryo-shaped pockets of polydimethylsiloxane (PDMS). Because of the optical transparency of materials used to construct the microfluidic device (glass and PDMS), embryos that have been labeled with fluorescent markers before trapping can be easily viewed under most microscope setups. Since the development of this microfluidic platform, we have used it in conjunction with immunostaining and

fluorescence in situ hydridization (FISH) to quantitatively study spatial and temporal patterning about the dorsal-ventral axis, and we have used the information to develop computational models that can predict these patterning events9–12.

Here we present a protocol that details the steps necessary for per-forming high-throughput, microfluidics-based imaging and data acquisition of dorsal-ventral gene patterning during Drosophila embryogenesis8 (Fig. 1). The microfluidic device can be used with most fluorescent microscopy setups, including upright and inverted epifluorescence confocal and multiphoton scopes, as well as super-resolution microscopy setups; it is operated with a single pumping source (e.g., syringe pump) for embryo loading onto the device. The chip consists of a single PDMS layer, which contains an inlet and outlet to a serpentine channel for embryo loading and cross-flow channels for automated loading and orientation of individual embryos within embryo traps (Fig. 1a). When the device is loaded under positive pressure (i.e., ~6 p.s.i.), the embryo traps expand owing to the mechanical deformability of PDMS, which allows embryos to enter the traps (Fig. 1b). After all traps are occupied with embryos, the pressure source can be turned off, returning the trap to a contracted shape to physically hold embryos in an end-on position. Once the device has been loaded with embryos, it can be mounted on a microscope for image acquisition. The ability and ease with which this device can be transported without loss of embryo orientation makes this device particularly useful in academic settings that possess public core facilities. For high- throughput image acquisition, it is necessary to integrate image-processing software such as MATLAB or LabVIEW with device operation13–15. The development of a robust image-processing program in this case is made simple, as embryos are immobilized at known positions within the device and in the same orientation.

Applications of the methodThe approach outlined in this protocol should be amenable to stud-ying other developmental events during D. melanogaster embryo-genesis that require spatial resolution about the dorsal-ventral

Microfluidic trap array for massively parallel imaging of Drosophila embryosThomas J Levario1, Mei Zhan2, Bomyi Lim3, Stanislav Y Shvartsman3 & Hang Lu1,2

1School of Chemical & Biomolecular Engineering, Georgia Institute of Technology, Atlanta, Georgia, USA. 2Interdisciplinary Program in Bioengineering, Institute of Biosciences and Bioengineering, Georgia Institute of Technology, Atlanta, Georgia, USA. 3Department of Chemical and Biological Engineering and Lewis-Sigler Institute for Integrative Genomics, Princeton University, Princeton, New Jersey, USA. Correspondence should be addressed to H.L. ([email protected]).

Published online 14 March 2013; doi:10.1038/nprot.2013.034

Here we describe a protocol for the fabrication and use of a microfluidic device to rapidly orient > 700 Drosophila embryos in parallel for end-on imaging. the protocol describes master microfabrication (~1 d), polydimethylsiloxane molding (few hours), system setup and device operation (few minutes) and imaging (depending on application). our microfluidics-based approach described here is one of the first to facilitate rapid orientation for end-on imaging, and it is a major breakthrough for quantitative studies on Drosophila embryogenesis. the operating principle of the embryo trap is based on passive hydrodynamics, and it does not require direct manipulation of embryos by the user; biologists following the protocol should be able to repeat these procedures. the compact design and fabrication materials used allow the device to be used with traditional microscopy setups and do not require specialized fixtures. Furthermore, with slight modification, this array can be applied to the handling of other model organisms and oblong objects.

http://www.nature.com/doifinder/10.1038/nprot.2013.034

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol

722 | VOL.8 NO.4 | 2013 | nature protocols

axis9–12,16–18. By scaling the dimensions of the embryo trap appropriately to the dimen-sion of the embryos, similar devices and the protocol should be capable of large-scale arraying of embryos in other Drosophila species. Furthermore, the hydrodynamic principles underlying the operation of the trap are applicable to biological speci-mens at other size scales, including mam-malian cells19, embryos of Caenorhabditis elegans and zebrafish (Danio rerio)20. We caution readers that the method of master fab-rication described in this protocol may not be the most appropriate for applications that require device heights above ~600 µm. Other methods such as silicon etching or macromachining techniques such as laser cutting should be evaluated in these scenarios.

We also envision this protocol incorporating other sophisticated techniques that have been used in previous work for probing bio-logical questions including, for example, laser microsurgery21,22. Laser-based technologies are readily accessible to the microfluidics- based approach outlined in this protocol, as the microscope glass slide part of the device is optically transparent to lasers used for microsurgery. Three-dimensional (3D), whole-organism imag-ing would also be an area in which we envision this device being applied, as one of the major drawbacks to 3D imaging is the embryo mounting23,24. The fact that over 700 embryos can be oriented in our device in less than 5 min would greatly increase the through-put of such studies. In addition to integrating more sophisticated techniques with the use of the embryo-trap array, we envision this device improving drug discovery studies involving Drosophila embryogenesis25,26. As the device accommodates a large number of embryos in a tightly packed viewing area, substantial improve-ments can be made in throughput and statistical reliability of such studies.

Comparison with other methodsCommon methods that have been used extensively for dorsal-ventral imaging in Drosophila before our microfluidics-based approach depended on image processing24,27 or manual manipulation28,29. Image reconstruction has been the primary method of image processing that was used to attain quantitative data about the spa-tial distributions of subcellular molecules around the dorsal-ventral axis. Liberman et al.27, for example, took confocal z-stack images of Drosophila embryos that were perpendicular to the dorsal-ventral plane; these were later combined with image processing to estimate the spatial distribution of Dorsal around the dorsal-ventral axis.

Keller et al.24, in contrast, took confocal z-stack images of Drosophila embryos from four perpendicular angles, and yet all images were still perpendicular to the dorsal-ventral plane. Both of these imaging modes suffer from limited spatial resolution within the dorsal-ventral plane relative to images that are taken while directly imaging the dorsal-ventral plane, and these images may include processing artifacts as both methods rely on image reconstruction programs. Furthermore, these methods of imaging the dorsal- ventral plane are throughput limited, as image reconstruction relies upon taking dense confocal z-stacks for high spatial resolution. For instance, although imaging in a direction parallel to the anterior-posterior axis requires at least ~150 z-stack images to reconstruct the dorsal-ventral plane, direct imaging along the anterior- posterior axis (i.e., in the dorsal-ventral plane) requires a single image to be taken, and thus it minimizes the optical and recon-struction artifacts.

Manual manipulation of embryos, as described by Witzberger et al.29, is a powerful method for immobilizing embryos in an end-on orientation (i.e., with the dorsal-ventral plane parallel to the cover glass). To fix embryos in an end-on orientation, Witzberger et al.29 machined a polyacrylamide gel such that it had cylindri-cal wells with dimensions comparable to those of a fly embryo (i.e., 400-µm height and 185-µm diameter). This platform was mounted on a cover glass and embryos were placed within the wells for end-on imaging. This platform was one of the first that could be used for end-on imaging with Drosophila embryos; however, the device still suffers from low throughput, because embryos were placed by hand into each well. In addition, the protocol used for making this device ultimately limited the packing density of the device, which could only hold 20 embryos. Our microfluidics-based approach to this problem is based on the technique developed by Witzberger et al.29, but improves upon it, as our design can rapidly load and orient > 700 embryos on one device in less than 5 min.

The major limitation of our technique is that it is currently only optimized for endpoint analysis or short-term observation. Although this is not a limitation for studies in which the use of

a c

Embryo

PDMS

PDMS

Cover glass

Objective

Cover glass

b

Figure 1 | Device and protocol overview. (a) The fabricated embryo-trap array microfluidic device loaded with green dye for visualization and the chip on a dissecting scope during embryo loading. (b) Schematic of the end-on orienting principle for imaging oblong fruit fly embryos along the dorsal-ventral axis. (c) Protocol sequence: microfabrication of SU-8 master (top), fabrication of PDMS molds (second from top), collecting and staining Drosophila embryos (second from bottom), and loading embryos onto the embryo-trap array for end-on imaging (bottom).

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


fixed embryos are sufficient for addressing the biological problem, this protocol is not currently recommended for studies requiring long-term observation of developmental dynamics. Although live embryos can be loaded and imaged in our device, optimal on-chip growth conditions have not yet been established. This may be due to oxygen limitations in such a densely packed array of live embryos. Furthermore, embryos currently cannot be recovered from the device after loading. Despite these limitations, the fabrication steps described in this protocol may still be useful for some live- imaging applications.

Experimental designDevice design and choice of fabrication technique. The operat-ing principle of our device is based on passive hydrodynamics, which allows automatic end-on orientation of Drosophila embryos during loading (Fig. 1b). Detailed design principles and a schematic of our device with dimensions are provided by Chung et al.8 and in Supplementary Figure 1. In brief, the design consists of a large ser-pentine channel and an array of cross-flow channels. The serpentine channel is used for bulk transport of suspended embryos onto the microfluidic device, whereas the cross-flow array creates a driving force (i.e., pressure drop) for embryo orientation and immobili-zation into the traps. Trapping requires optimization of channel dimensions and flow rate, but end-on orientation is solely depend-ent upon the geometry of the embryo and trap design. In our case, D. melanogaster embryos are ellipsoidal in shape with a long axis of ~500 µm and a short axis of ~150 µm. We designed the traps to be a truncated cylinder with a height of 500 µm and a diameter of 150 µm, because this structure is easily fabricated using the stand-ard lithographic techniques (Fig. 1b). Depending on the size and shape of the embryos of interest, one can easily modify the design accordingly. Our device without modifications should be compat-ible with most Drosophila species as the size and shape of embryos of these species are fairly similar. However, scaling the design up slightly (i.e., by increasing the trap height and diameter) for the slightly larger D. sechellia embryos would likely result in better trapping30. Scaling up of our device design for other model organ-isms should be straightforward and it has been done with zebrafish embryos20. It is noteworthy that for the zebrafish embryos, however, there was no orientation as they are nearly spherical and our design relies on the asymmetry in the embryo shape to achieve orientation. As noted above, master fabrication may also require alternative methods for larger embryos ( > 600 µm). We also note that masters may also be fabricated using foundry services such as the Stanford Microfluidic Foundry (http://www.stanford.edu/group/foundry/) with the caveat that more stringent geometric constraints (such as aspect ratios) may be imposed by the manufacturers.

In order to pick the material with which the final device is made, one needs to consider a few things. First, one of our design con-straints is the use of a flexible material, as our design requires the device to deform and expand slightly during loading, and it contracts to hold the embryos for imaging in end-on orienta-tion. PDMS is an ideal choice. In addition to its elasticity, PDMS is optically transparent in the visible spectrum and is low in auto-fluorescence. Furthermore, PDMS is easily bonded to glass slides or cover glasses through various procedures such as plasma treatment, which lends it more useful to microscopic investigation. PDMS is thus generally considered one of the better materials for making our microfluidic device.

Constructing PDMS devices is accomplished by replica mold-ing from a master. Masters can be fabricated using surface or bulk micromachining or conventional machining. Bulk or surface micromachining silicon wafers are possible, but processes either do not allow for nonsloped walls or require expensive equipment not widely available. Many of the processes are also too slow or too variable. Finally, most standard 100-mm silicon wafers are 500 µm thick, a thickness that is similar to most device thicknesses and would require more sophisticated steps such as wafer bonding.

In comparison, photolithographic technique with thick resists is much more straightforward and has a fast turnaround time. Typically, data from manufacturers of photoresists are sufficient for optimizing the process to make masters with the appropriate height. In-plane features are determined by the masks and thus can be well controlled. Photolithography also requires (fewer pieces of) simpler equipment than silicon etching, as only a spin coater and a mask aligner are needed. Microchem SU-8 negative epoxy series resists are an ideal choice as they allow high-aspect-ratio structures that are very thick (up to 600 µm in a single coat), although other thick resists are also possible.

Embryo preparation considerations. Our device is compatible with any method for preparing embryos to image Drosophila embryogenesis. This compatibility results from the fact that embryos are prepared off-chip before embryo loading and the device does not alter the markers of interest. Embryos can be fluo-rescently labeled by either endogenous expression (i.e., transgenes) or exogenous staining (e.g., immunostaining). Immunostaining was the preferred method used in our previous work as it is a much simpler technique for imaging multiple proteins of inter-est in a single embryo8. Successfully imaging multiple proteins of interest, however, does require well-selected fluorophores (e.g., nonoverlapping emission and excitation wavelengths) and antibodies (if performing immunostaining). However, these are issues that are common to all fluorescence-based imaging tech-niques. Our system is not limited to imaging just proteins, and it can be compatible with methods such as FISH9,31 for imaging DNA and RNA transcripts.

Loading and microscopic considerations. Loading embryos onto our device is simply accomplished by pumping embryos in suspen-sion using a pressure source. We chose to use buffers such as PBS with trace amounts of a surfactant such as Triton X-100 or BSA. The surfactant or BSA is used to prevent embryo aggregation and clogging in the microchannels and tubing. The surfactant should be compatible with the embryos and the fluorophores used for labe-ling proteins or nucleic acids of interest. PBS is used because it has a low viscosity and requires a low pressure to flow into the device. The buffer, however, should be exchanged with a buffer containing glycerol for fixed embryos, thereby matching the refractive index of the PDMS and enhancing the image quality. It is important to know what magnification a particular application will require and consequently what the working distance of the objective will allow. The working distance will dictate how thick a cover glass can be for imaging.

Microscopy setups are more dependent on the particular application and are not limited by our device. If one so chooses, two separate microscopes can be used for loading and imaging. Transporting a loaded device from one microscope to another is

http://www.stanford.edu/group/foundry/

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


allowed by the embryo trap design, which gently yet securely holds embryos in an end-on position. The use of two microscopes has the following advantages. During loading, a large viewing area is desired so that all or most of the device can be monitored for load-ing errors (e.g., clogging), which can be addressed appropriately. A large-enough viewing area for this massively parallel imaging device is best provided by a dissecting scope or stereomicroscope, as these have large working distances and allow manual intervention if needed. During imaging, high magnification and high resolu-tion are usually desired; for larger specimens, such as Drosophila

embryos, this typically requires the use of a confocal microscope. As PDMS is transparent in the visible wavelengths, imaging can also be conducted in a variety of modes, including transmitted light and reflected light; however, we recommend for quantitative analysis that embryos be imaged through the cover glass. Furthermore, the robust immobilization that the microfluidic device provides also allows inverting the device so that it can be imaged through the cover glass in either an upright or inverted microscope. The flex-ibility of our system allows many choices to be made by the user for his or her particular application.

MaterIalsREAGENTSGeneral reagents

Distilled waterIsopropanol, 4 liters (Sigma-Aldrich, cat. no. 437522) ! cautIon Isopropanol is highly flammable. Ensure that you wear proper personal protective equipment (PPE), keep away from ignition sources and use isopropanol within a fume hood.DPBS (Dulbecco’s PBS, VWR, cat. no. 45000-436)Stained embryos (or adult flies for collecting and staining embryos in house; see embryo immunostaining, Box 1)

Microfabrication of SU-8 masterSulfuric acid, 500 ml (Sigma-Aldrich, cat. no. 320501) ! cautIon Sulfuric acid is corrosive. Ensure that you wear proper PPE, which includes a face shield, acid-proof gown and acid-proof gloves, and use this reagent within a fume hood.Hydrogen peroxide, 100 ml (Sigma-Aldrich, cat. no. 216763) ! cautIon Hydrogen peroxide is toxic. Ensure that you wear proper PPE and use this reagent within a fume hood.SU-8 2010 negative photoresist (Microchem) ! cautIon Fumes from photoresist are toxic. Ensure that you wear proper PPE, and use this reagent within a fume hood.SU-8 developer (propylene glycol monomethyl ether acetate; Eastman) ! cautIon Propylene glycol monomethyl ether acetate is toxic. Ensure that you wear proper PPE, and use it within a fume hood.Silanizing agent ((tridecafluoro-1,1,2,2-tetrahydrooctyl)-1-trichlorosilane; United Chemical Technologies, cat. no. T2492) ! cautIon This reagent is toxic. Ensure that you wear proper PPE and work in a fume hood.Piranha solution

Fabrication of PDMS devicePDMS (Sylgard 184 silicone elastomer kit; Dow Corning)

Embryo immunostainingBleach, 25 ml (Sigma-Aldrich, cat. no. 239305) ! cautIon Bleach is toxic and corrosive. Ensure that you wear proper PPE and work in a fume hood.Grape (or apple) juice agar plate (Reagent Setup)Agar (Genesee Scientific, cat. no. 66-103)Methyl paraben (Sigma-Aldrich, cat. no. 47889)Ethanol, 500 ml (Sigma-Aldrich, cat. no. E7148) ! cautIon Ethanol is highly flammable. Ensure that you wear proper PPE, keep away from igni-tion source and use this reagent within a fume hood.Grape (or apple) juice concentrateYeast, 500 g (Sigma-Aldrich, cat. no. YSC2)Formaldehyde solution, 100 ml (Sigma-Aldrich, cat. no. 252549) ! cautIon Formaldehyde is toxic and flammable. Ensure that you wear proper PPE, keep away from ignition source and work in a fume hood.Heptane, 1 liter (Sigma-Aldrich, cat. no. 494526) ! cautIon Heptane is highly flammable. Ensure that you wear proper PPE, keep away from ignition source and use it within a fume hood.Triton X-100 surfactant, 100 ml (Sigma-Aldrich, cat. no. X100-100ML)Methanol (Sigma-Aldrich, cat. no. 154903-2L) ! cautIon Methanol is highly flammable. Ensure that you wear proper PPE, keep away from ignition source and use this reagent within a fume hood.Image-iT signal enhancer (Invitrogen, cat. no. I36933)BSA solution, 50 ml (BSAS; Sigma-Aldrich, cat. no. A7284)Normal whole goat serum (NGS; VWR, cat. no. RLB304)

••

••

•

•

•

•

•

•

•

•

••••

•••

•

••

•••

Primary antibody for your experimental needs (Invitrogen)Secondary antibody for your experimental needs (Invitrogen)Alexa Fluor dye for your experimental needs (Invitrogen)PBS with Triton X-100 (PBST, Reagent Setup)

Device operationGlycerol, 100 ml (Sigma-Aldrich, cat. no. G5516)Powdered BSA, 50 g (BSAP; Sigma-Aldrich, cat. no. A4503)PBS with BSAP (PBSB, Reagent Setup)

EQUIPMENTGeneral equipment

Tape (Scotch Magic tape)Razor blade (McMaster-Carr, cat. no. 3962A1)LighterPetri dish, 100 and 150 mm (VWR, cat. nos. 25384-302 and 25384-326)Pipettors, 0.5–10, 10–100 and 100–1,000 µl (VWR)Pipette tips, 0.1–10, 1–200 and 100–1,250 µl (VWR)Dissecting scope (Top illumination is necessary for viewing the wafer. Bottom illumination is preferred for device loading; Zeiss)Needle, 19- and 20-gauge (1/2-inch Luer needle, blunt tip, 19 and 20 gauge; VWR)Vacuum desiccator (Fisher Scientific, cat. no. 420250000) and vacuum pump (Thermo Scientific, cat. no. 420–1901)Kimwipes (VWR, cat. no. 21905-011)Alcohol burner (VWR, cat. no. 17805-005)Conical tubes

Microfabrication of SU-8 masterSilicon wafer (100 mm; University Wafer)Aluminum top hot plate (65, 95 and 200 °C; McMaster-Carr, cat. no. 3271K14)Spin coater with vacuum wafer chuck for 4-inch wafers (Laurell)Mask aligner with near-UV exposure (360 nm; SUSS MicroTec) ! cautIon UV light can burn eyes. Use protective eyewear.Intensity meter (SUSS MicroTec)Teflon bowl (SPI, cat. no. 01730-AB)Glass crystallizing dish (VWR, cat. no. 89090-662)Wafer rack (Teflon)Stir bar (VWR)Stir plate (VWR)Photomask (dark field, embryo-trap design; CAD Art Services)Glass filter (127 mm × 127 mm, 360-nm long-pass mask aligner filter; Omega Optical, cat. no. 2007308)Level (McMaster-Carr)Wafer tweezers (SPI, cat. no. 0S4WF-XD)Aluminum foilPPE (including acid-resistant gloves, acid-resistant gowns and acid-resistant face shields)Amber bottle (VWR, cat. no. 16180-728)Nitrogen gun

Fabrication of PDMS deviceBalance (Fisher Scientific, cat. no. S94792B)Oven (Fisher Scientific, cat. no. 13-247-625G)Disposable cups (150 ml; VWR, cat. no. 15704-010)Disposable spatula (VWR, cat. no. 80081-188)Rotary tool (McMaster-Carr, cat. no. 53625A44) with drum sander tip (McMaster-Carr, cat. no. 4648A13) or a flat file for sharpening needles

••••

•••

•••••••

•

•

•••

••••

••••••••

••••

••

•••••

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


Box 1 | Embryo immunostaining ● tIMInG ~7 h plus overnight incubation collection of stage 4 embryos 1. Place adult D. melanogaster (~50 flies) in an embryo collection cage with a grape juice agar plate.2. Place a large coin-sized circle of inactive yeast in the center of the grape juice agar plate (food for adult flies during collection). Cover the plate with a box to simulate night. Let it sit for 3 h (a 3-h stretch of time gives time for flies to lay eggs in the agar plate and develop so that the majority of embryos collected will be at stage 4 of embryogenesis).3. Remove the agar plate from the embryo collection cage and discard the adult flies. Embryos on the agar plate are easily visible with the naked eye.4. Remove the inactive yeast by gently scraping the agar with a plastic spoon. Remove any dead flies from the agar plate using forceps.

chorion removal5. Assemble the nylon mesh into a conical shape and clip it into place using forceps. Spray the mesh with distilled water before pouring embryos.6. Pour a ~2.5% (wt/vol) sodium hypochlorite solution (typically a 1:1 mixture of bleach:water) onto the agar plate. Use a plastic spoon to gently release embryos from the agar surface. Agitate the plate with a spoon for 1.5 min.7. Strain the embryos using the nylon mesh and rinse them with distilled water, removing any excess bleach, for at least 1 min. crItIcal step Ensure that you prepare the fixation solution before dechorionation.

embryo fixation8. Prepare the fixation solution (do so before step 5 in Box 1) by pooling together 4 ml of PBS and 5 ml of heptane in a 20-ml scintillation vial.! cautIon Heptane is highly flammable. Keep it away from ignition sources and perform this step in a fume hood.9. Place embryos in the fixation solution and add 1 ml of 37% (wt/vol) formaldehyde. Gently mix on a rocker for 20–25 min. Two phases should be visible (upper = heptane and lower = PBS + formaldehyde) and the embryos should be located at the interface.! cautIon Formaldehyde is toxic and flammable. Proper PPE should be used; this step should be performed in a fume hood and away from ignition sources. crItIcal step Timing is important. Formaldehyde must be placed in the fixation solution in less than 1 min after adding the embryos.

removal of the vitelline membrane (devitellinization)10. Remove the lower aqueous phase (i.e., the PBS and formaldehyde) using a 100–1,000-µl pipettor. Add 5 ml of methanol and shake vigorously for 75 s. Devitellinzed embryos will sink to the bottom of the vial, whereas non devitellinized embryos and debris will remain at interface.11. Remove the upper heptane phase with the pipettor. Add 5 ml of methanol to the scintillation vial and shake it vigorously again for 15 s.12. Remove embryos that have collected at the bottom of the scintillation vial using a 100–1,000-µl pipettor and place them in a 1.5-ml Eppendorf tube.13. Wash the embryos three times by adding 1 ml of methanol, gently shaking and removing the methanol. Add 1 ml of methanol. pause poInt The embryos thus treated can be immunostained or stored at − 20 °C for later use. Embryos can be stored for up to 1 month.

embryo immunostaining14. Remove the methanol from the 1.5-ml Eppendorf tube that contains fixed embryos with a 100–1,000-µl pipettor and rehydrate embryos with PBST. Make the transition from methanol to PBST gradual over a few washes with mixtures of PBST and methanol that increase with respect to the ratio of PBST to methanol. crItIcal step In this and the subsequent steps,make sure to protect the Eppendorf tube from exposure to light.15. Wash embryos twice by adding 1 ml of PBST to the Eppendorf tube with the fixed embryos and gently mixing on a rocker for 5 min. Remove as much of the washing solution as possible.16. Add Image-iT signal enhancer by adding four or five drops of Image-iT signal, and then gently mix for 30 min on a rocker. Remove the Image-iT with a pipettor. Wash embryos twice by adding 1 ml of PBST and by gently mixing for 5 min each on a rocker. Remove PBST with a pipettor.17. Block for 90 min at room temperature in 1 ml of a mixture of 50 µl of NGS, 300 µl of BSAS and 650 µl of PBST.18. Attach the primary antibody by incubating overnight at 4 °C in 0.5 ml of the first antibody solution, which contains 1:100 ratio of the primary antibody, 25 µl of NGS, 15 µl of BSAS and 650 µl of PBST. Rinse the embryos twice in 1 ml of PBST. Rinse further by adding 1 ml of PBST to the Eppendorf tube and gently mixing for 1 h on a rocker while changing the solution twice throughout.19. Attach the secondary antibody by incubating the embryos for 90 min at room temperature in 0.5 ml of the second antibody solution, which consists of 1:500 Alexa Fluor dye, 50 µl of NGS, 15 µl of BSAS and 650 µl of PBST. If you want to visualize nuclei, use DAPI in this step.20. Rinse two times in 1 ml of PBST by closing the tube, flipping the tube five times and removing the PBST. Rinse further by adding 1 ml of PBST to the Eppendorf tube and gently mix the tube for 1 h while changing the solution twice throughout.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


Cover glass (VWR, cat. no. 48393-252) or glass slide (VWR, cat. no. 16004-430)Plasma chamber (Harrick, cat. no. PDC-32G) and vacuum pump (Robinair/CoolTech, model 15600) crItIcal If you are not using this particular plasma chamber and vacuum pump, then plasma bonding will require optimization of step timing, especially for the PDMS exposure time to the plasma environment.Compressed air source (e.g., house air)Scalpel with a no. 11 scalpel blade (McMaster-Carr, cat. no. 36325A63 and 36325A71)

Embryo immunostainingGentle rocker (Fisher Scientific, cat. no. 260100F)Disposable spoonNylon mesh (McMaster-Carr, cat. no. 9318T23)Forceps (VWR)1.5-ml Eppendorf tubes (VWR, cat. no. 20170-650)Embryo collection cage (Genesee Scientific, cat. no. 59-101)20-ml scintillation vial (VWR, cat. no. 66022-106)

Device operation and imagingInlet reservoir (2×, Equipment Setup)Connection pin (2×, Equipment Setup)Pressure control system (Equipment Setup)Needle-nose pliers (McMaster-Carr)Adjustable wrench (2×; McMaster-Carr)Camera (Lumenera) with computer connectivity and compatible computerCentrifuge tube, 50 ml (2×; VWR, cat. no. 21008-940)Centrifuge tube rack (VWR, cat. no. 60916-101)Teflon tape (McMaster-Carr, cat. no. 4591K11)Epoxy (J-B Weld original cold weld epoxy system)Manual pinch valves for 3/32-inch outer diameter (o.d.) tubing (McMaster-Carr, cat. no. 5031K11)A bin for collecting waste from the microfluidic device outlet (e.g., 250 ml flask)Silicone tubing (1/32-inch inner diameter (i.d.) × 3/3-inch o.d., ~5 feet; McMaster-Carr, cat. no. 5236K501)Regulator (pressure regulator for air, 10–32 UNF threaded, 0.5–30 p.s.i.; McMaster-Carr, cat. no. 43275K13)Pressure gauge (1/8-inch NPT input adapter, 0–15 p.s.i.; McMaster-Carr, cat. no. 3846K41)Push-to-connect Y junction (5/32-inch o.d. tube, maximum pressure > 150 p.s.i.; McMaster-Carr, cat. no. 5111K402)Nylon tubing (Extra-flex nylon tubing .106-inch i.d., 5/32-inch o.d., .025-inch wall thickness, ~5 feet; McMaster-Carr, cat. no. 5112K52)Pressure gauge adapter (push-to-connect adapter for 5/32-inch tube o.d. × 1/8-inch NPT female pipe; McMaster-Carr, cat. no. 5111K664)Regulator adapter (3×, push-to-connect adapter for 5/32-inch tube o.d. × 10–32 UNF male thread; McMaster-Carr, cat. no. 52065K124)Coupler (10–32 thread size, through-wall coupling; McMaster-Carr, cat. no. 5454K85)Luer lock adapter (male Luer integral lock ring × 10–32 special tapered thread; Value Plastics, cat. no. XMTLL-6)Stopcock (FDA Luer lock stopcock, male × male; McMaster-Carr, cat. no. 7033T11)Pressure source adapter (push-to-connect adapter for 5/32-inch tube o.d. × appropriately sized fitting for pressure source outlet, see Equipment Setup; McMaster-Carr)

REAGENT SETUPGrape juice agar plate As previously published in ref. 32. In brief, dissolve 25 g of agar in 700 ml of water via autoclaving for 40 min. Dissolve 0.5 g of methyl paraben in 20 ml of 95% (vol/vol) ethanol and add to 300 ml of grape (or apple) juice concentrate. Add the grape juice mixture to the autoclaved agar and pour it into 100-mm Petri dishes (this size fits into the large embryo collection cage). Plates can be made well in advance (~1 month) and stored at 5 °C.PBST Mix 498.5 ml of DPBS with 1.5 ml of Triton X-100. This can be made well in advance (~1 month) and stored at room temperature (22 °C).PBSB Mix 5 g of BSAP with 500 ml of PBS to create 1% (wt/vol) PBSB. This can be made well in advance (~1 month) and stored at 5 °C.EQUIPMENT SETUPSharpened 19-gauge blunt-tip needle Sharpen a 19-gauge blunt-tip needle with an electric rotary tool with a drum sander attachment. In order to sharpen, sand the tip of the needle at a 45° angle with a drum sander.

••

••

•••••••

•••••••••••

•

•

•

•

•

•

•

•

•

•

•

•

Sharpen and clean the interior surface of the needle by running a no. 11 scalpel blade inside the needle. Test the needle on waste pieces of PDMS, ensuring that it creates a smooth hole before using it on devices.Making connection pins Make six connection pins as follows: grasp the metal end of the 1/2-inch Luer, blunt-tip, 19-gauge needle with needle-nose pliers. Quickly brush the Luer end of the needle over a flame from the alcohol burner (~5 s). Grab the Luer end of the needle and pull to dislodge it from the metal end (Fig. 2). When removing the Luer end, ensure that you do not grasp the metal needle too tightly with the pliers as it may bend the metal tip, which is undesired. Typically, the Luer end will not be removed cleanly and leave some residual plastic on the metal tip. Clean the metal tip by again brushing the plastic over the flame (~5 s) and wipe it clean with a Kimwipe. Keep the now-bare metal end of the needle—this is the connection pin. ! cautIon The Luer needle will be hot; ensure that you protect your hands with some insulation such as a Kimwipe.Inlet reservoir Construct two reservoirs with the following steps: punch two holes on the top of a 50-ml centrifuge tube cap using a red hot 1/2-inch Luer, blunt-tip, 20-gauge needle (red hot from holding over alcohol burner for ~5 s) (Fig. 2b). Insert two of the connection pins into the holes so that one half of the pin is above the cap and the other half is below the cap. Prepare the J-B Weld original cold weld epoxy by thoroughly mixing the two tubes of fluid in a 1:1 (vol/vol) ratio. This is most easily accomplished by pouring a pea-sized droplet from each tube into a Petri dish and stirring with a pipette tip. Apply the epoxy at the needle-cap interface on both sides of the cap to create an air-tight seal. Allow to cure undisturbed overnight. Attach a length of silicone tubing to the inside of one of the connection pins so that it reaches near the bottom of a conical tube when the cap is screwed on (~10 cm). This will be the pin that needs to be connected to the microfluidic device (Fig. 2c,d). Make two of the inlet reservoirs. ! cautIon Inlet reservoirs should be constructed at least 1 d in advance of a device-loading experiment, thereby allowing epoxy to thoroughly cure at room temperature. Reservoirs can be used multiple times as long as the epoxy seals the cap. crItIcal This needs to be done at least 1 d in advance to allow the epoxy to completely cure before use.Pressure control system The pressure control system that is attached to the pressure source is constructed with the following steps (Fig. 2e,f). Use an appropriate adaptor for converting the output of the compressed air source to a 5/32-inch o.d. push-to-connect connector. The adaptor will depend on the specific compressor or house air connection in use. Most commonly, the outlet will be a female pipe connector. In this case, select an appropriate 5/32-inch push-to-connect male pipe adapter to mate to the female pipe thread. Attach the 5/32-inch tube o.d. × 1/8-inch NPT female pipe adapter to the pressure gauge. Secure tightly using two wrenches. Attach two of the 5/32-inch tube × 10–32 UNF male thread adapters to the regulator. Secure tightly with wrench. The particular adapters listed in the materials list have a rubber washer, and thus application of Teflon tape is not necessary. Attach a 5/32-inch tube × 10–32 UNF male thread adapter to the 10–32 thread size coupler. Secure tightly with two wrenches. Attach the male Luer × 10–32 thread adapter to the other end of the coupler. Attach a 1/2-inch Luer, blunt-tip, 19-gauge needle to the Luer adapter. Attach two 1/2-inch Luer, blunt-tip, 19-gauge needles to the male × male, Luer lock stopcock. Connect the pressure source adapter to the regulator inlet with nylon tubing (5/32-inch o.d.). Typically, the inlet and outlet ports for the regulator will be specified on the regulator. Connect the regulator outlet to the base of the Y-junction (5/32-inch o.d.) with nylon tubing. Connect one of the Y-junction outlets to the pressure gauge adapter with nylon tubing. Connect the other Y-junction outlet to the 5/32-inch tube × 10–32 UNF male thread adapter with nylon tubing. Connect the Luer needle from the coupler to one of the Luer needles on the stopcock with silicone tubing. crItIcal Wrap a single layer of Teflon tape around the male pipe threading on the pressure gauge before connecting the female pipe adapter to the pressure source. This will help in providing air-tight seals between the connections. crItIcal We recommend that first-time builders use the specific pieces outlined in this protocol, as the design of the pressure control system will require that all pieces are compatible with each other (i.e., size). However, the particular pieces described here are not necessarily required. If one chooses to construct a custom system, it is necessary to use parts that are rated to withstand pressures for which the pressure source is rated.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


proceDureMicrofabrication of su-8 master ● tIMInG ~5 h plus overnight silanization1| Piranha clean. New wafers should be piranha cleaned before use (Fig. 3). Place a glass crystallizing dish (150 mm diameter × 75 mm height) with 100 ml of sulfuric acid on a 125 °C hot plate. Cover the hot plate with aluminum foil to protect it from the piranha solution. Once sulfuric acid is heated up to the desired temperature, add ~33 ml of hydrogen peroxide. Place the wafer in the piranha solution for 10 min. Remove and rinse thoroughly with distilled water. Dry with nitrogen gun.! cautIon Sulfuric acid is corrosive. Ensure that you wear proper PPE, which includes a face shield, acid-proof gown and acid-proof gloves. Perform this inside a fume hood. Ensure that you use wafer tweezers that are compatible with piranha solution when retrieving the wafers from the bath. crItIcal step The mixture will bubble if it is at the correct temperature of 125 °C.

2| Before working on the hot plates and spin coaters, which will be used during film deposition, soft baking and postexposure baking, ensure that you level the surfaces where the wafer will be placed (i.e., the hot plates and spin coater vacuum chucks). crItIcal step It is very important that these areas be leveled, because the SU-8 film is still fluid and deformable during baking.

3| Dehydrate. Place a piranha-cleaned silicon wafer polished side up on a 200 °C hot plate for 15 min. Remove it using a wafer tweezer and allow the wafer to cool for ~5 min.

4| Adhesion-layer deposition. Line the spin coater with an aluminum foil to catch excess SU-8. Place the wafer on the vacuum chuck within the spin coater while ensuring that the wafer is centered on the vacuum chuck. Dispense ~1 ml of SU-8 2010 at the center of the wafer and run a two-step program with the following settings. First, use a spin rate of 500 r.p.m. for 5 s with an acceleration of 100 r.p.m. per second; second, use a spin rate of 3,000 r.p.m. for 30 s with an acceleration of 300 r.p.m. per second (Fig. 3b). Once the program has been completed, remove the wafer from the spin coater.! cautIon SU-8 photoresists are toxic. Ensure that you wear proper PPE, and perform this inside a fume hood. crItIcal step When pouring any SU-8 series photoresist, it is important to pour as close to the wafer surface to prevent air bubbles from getting trapped in the SU-8 photoresist. Air bubbles are not easily removed from SU-8 because of its high viscosity, and they will persist in the film even after spinning, thereby ruining the uniformity and physical integrity of the film. It is suggested that SU-8 photoresists be transferred to a small, wide-mouth amber glass bottle. This will allow close and controlled pouring of SU-8 photoresists. crItIcal step Centering the wafer is important for uniform film deposition of any photoresist and secure immobilization on the vacuum chuck. Wafers that are not correctly centered tend to dislodge from the vacuum chuck during spinning, thereby ruining the film and possibly breaking the silicon wafer. Spin coaters typically have a button that can test the wafer centering; this basically spins the wafer so the user can visualize whether the wafer is centered. If the spin coater does not

a b c

d e f

g h i

Figure 2 | Equipment setup. (a) Removing the plastic Luer stub from a 19-gauge needle to leave behind the hollow metal pin (connection pin). (b) Punching holes in the cap of a 45-ml conical tube with a hot 20-gauge needle for the reservoir. (c,d) The assembled cap of the reservoir with 19-gauge connection pins fixed into the cap with epoxy and a length of silicone tubing. (e) The parts for the pressure control system laid out according to connectivity. (f) Fully assembled pressure control system (adapter for pressure source located at the far right). (g) The reservoir, pressure control system, PDMS device, connecting tubing and pins, manual pinch valves and waster flask before connection. (h) The connection pins placed into the inlet/outlet of the PDMS device with silicone tubing. (i) The fully assembled system.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


have this option, then a dry run can be performed without any photoresist in order to visually inspect the wafer centering. If the wafer is not centered, adjust it and test again. crItIcal step When transporting coated wafers over large distances, we recommend placing the wafer inside a 150-mm Petri dish to protect the surface from particulates such as dust.? trouBlesHootInG

5| Soft bake. Place the wafer on a 95 °C hot plate for 3 min. Remove the wafer from the hot plate and allow the wafer to cool for ~5 min. It is suggested to grasp the silicon wafer with the wafer tweezers in the same area every time. Wafer tweezers will ruin the deposited film where they touch the wafer, so grasping a single spot will minimize the loss of usable area on the wafer. Film should appear smooth with no defects (e.g., cracking, Fig. 4).! cautIon Fumes from photoresist are toxic. Perform this step inside a fume hood.

6| Exposure. Place the glass filter into the mask holder of the mask aligner. Place the wafer on the base of a mask aligner. Expose the entire wafer to 365-nm light with an energy dose of 100 mJ cm − 2. crItIcal step Energy dose is determined by the intensity of the light and exposure time. Ensure that you measure the light intensity ([ = ] mW cm − 2) through the 360-nm filter before loading the wafer and mask into the mask aligner. Dividing the energy dose by the light intensity gives the exposure time in units of seconds.? trouBlesHootInG

a b c d

e f g h

i j k l

Figure 3 | Microfabrication of SU-8 master. (a) Piranha cleaning. (b) Pouring SU-8 photoresist onto the wafer in the spinner lined with aluminum foil. (c) Coated silicon wafer with 10-µm-thick adhesion layer. (d) The silicon wafer after three cycles of spinning with SU-8 2100 (thickness ~500 µm). (e) The photomask taped to the 127 × 127 mm 365-nm glass filter in the mask holder of a mask aligner. (f) The patterned wafer after exposure; features are visibly present after an energy dose of 560 mJ cm − 2. (g) The patterned wafer after the postexpose bake; features are more prominent. (h) The development setup with a stir bar inside a Teflon wafer boat and the silicon wafer at position 4 in the wafer boat with the feature side down. (i) The patterned wafer within the SU-8 developer during development. (j) An underdeveloped wafer with clearly visible undeveloped SU-8 still present on the wafer surface. (k,l) A fully developed wafer (k) that is ready for silanization (l) within the vacuum desiccator.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


7| Postexpose bake. Place wafer on a 95 °C hot plate for 3 min. Remove the wafer from the hot plate and allow the wafer to cool for ~5 min (Fig. 3c). Film should appear smooth with no defects.! cautIon Fumes from photoresist are toxic. Perform this inside a fume hood.

8| Feature-layer deposition. Line the spin coater with aluminum foil to catch excess SU-8. Place the wafer on the vacuum chuck within the spin coater while ensuring that the wafer is centered on the vacuum chuck. Dispense ~1 ml of SU-8 2100 at the center of the wafer and run a two-step program with the following settings. First, use a spin rate of 500 r.p.m. for 5 s with an acceleration of 100 r.p.m. per second; second, use a spin rate of 1,000 r.p.m. for 30 s with an acceleration of 300 r.p.m. per second (Fig. 3b). Once the program has been completed, remove the wafer from the spin coater.! cautIon SU-8 photoresists are toxic. Ensure that you wear proper PPE, and perform this inside a fume hood. crItIcal step When pouring any SU-8 series photoresist, it is important to pour as close to the wafer surface to prevent air bubbles from getting trapped in the SU-8 photoresist. Air bubbles are not easily removed from SU-8 because of its high viscosity; air bubbles will persist in the film even after spinning, thereby ruining the uniformity and physical integrity of the film. It is suggested that SU-8 photoresists be transferred to small, wide-mouth amber glass bottle. This will allow close and controlled pouring of SU-8 photoresists. crItIcal step Centering the wafer is important for uniform film deposition of any photoresist and secure immobilization on the vacuum chuck. Wafers that are not correctly centered tend to dislodge from the vacuum chuck during spinning, thereby ruining the film and possibly breaking the silicon wafer. Spin coaters typically have a button that can test the wafer centering; this basically spins the wafer so the user can visualize whether the wafer is centered. If the spin coater does not have this option, then a dry run can be performed without any photoresist in order to visually inspect the wafer centering. If the wafer is not centered, adjust it and test again. crItIcal step When transporting coated wafers over large distances, we recommend placing the wafer inside a 150-mm Petri dish to protect the surface from particulates such as dust.? trouBlesHootInG

9| Soft bake. Place the wafer on a 65 °C hot plate for 7 min, and immediately follow that by placing the wafer on a 95 °C hot plate for 45 min. Remove the wafer from the hot plate and allow it to cool for ~5 min. Film should appear smooth with no defects.! cautIon Fumes from photoresist are toxic. Perform this inside a fume hood.

10| Repeat Steps 8 and 9 for a total of two feature-layer depositions and two soft bakes. This layering of two films on the top of each other produces the desired thickness of 500 µm for the feature layer (Fig. 3d). (Each round of feature layer depositions deposits a film of ~250 µm thick.)

11| Exposure. Assemble the photomask and glass filter by taping the photomask to the glass filter (Fig. 3e). Place the photomask–glass filter into the mask holder of the mask aligner such that when the mask and the wafer come into contact, the photomask will contact the wafer surface. Place the wafer on the base of a mask aligner. Expose the entire wafer to 365-nm light with an energy dose of 560 mJ cm − 2. Features should become immediately visible as a result of the high energy dose required for the thick film of SU-8 (Fig. 3f) (this is generally an indicator of overdosing in thin films, as stated in the user guidelines of SU-8 products). crItIcal step Energy dose is determined by the intensity of the light and exposure time. Ensure that you measure the light intensity ([ = ] mW cm − 2) through the 360-nm filter before loading the wafer and the mask into the mask aligner. Dividing the energy dose by the light intensity gives the exposure time in units of seconds.? trouBlesHootInG

a

b

c

d

Figure 4 | Common problems. (a) Cracking of SU-8 film during the soft or postexpose bake. (b) Delaminating during development. (c) Delaminating during the peeling of PDMS molds (e.g., missing cross-flow channels and embryo traps) and/or when PDMS molds are not cleanly removed during peel (right, in between focusing channels). Scale bar, ~1 mm. (d) Embryo clogging during loading. Scale bar, ~800 µm.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


12| Postexpose bake. Place the wafer on a 65 °C hot plate for 5 min, and immediately follow that with placing the wafer on a 95 °C hot plate for 12 min. Remove the wafer from the hot plate and allow to cool for 5 min. Features should become more prominent after the postexpose bake (Fig. 3g).! cautIon Fumes from photoresist are toxic. Perform this inside a fume hood.? trouBlesHootInG

13| Development. Develop the wafer in the SU-8 developer (propylene glycol monomethyl ether acetate) for ~25–30 min. Continuous agitation is most easily accomplished by placing a Teflon wafer rack on its end with a stir bar inside the rack and the wafer at the fourth position above the stir bar with the film side down (Fig. 3h). Place the setup inside a bowl containing the SU-8 developer on top of a stir plate and continue developing for the required time (Fig. 3i).! cautIon Propylene glycol monomethyl ether acetate is toxic. Ensure that you wear proper PPE, and perform this inside a fume hood. crItIcal step On removing the wafer from the developer, it is important ensuring that the unexposed SU-8 was completely developed. Undeveloped SU-8 can be easily seen when the wafer is severely underdeveloped (Fig. 3j). Undeveloped SU-8 may not necessarily be visible even after longer development time, but it can be checked for by spraying the wafer with isopropyl alcohol. If there appears to be a white colloidal suspension in the isopropyl alcohol, then there is still undeveloped SU-8 present, which must be removed by placing the wafer back into the developer for ~5 min more. Continue developing the wafer and checking with isopropyl alcohol until the wafer is fully developed (Fig. 3k). SU-8 structures should remain attached to the wafer throughout development; they should not, for example, delaminate (Fig. 4b).? trouBlesHootInG

14| Rinse the wafer with isopropyl alcohol and thoroughly dry the wafer with a nitrogen gun.

15| Silanization. Place the wafer inside a desiccator with a small vial of the silanizing agent (~1 ml, (tridecafluoro-1,1,2, 2-tetrahydrooctyl)-1-trichlorosilane); and pump it down for ~5 min (Fig. 3l). Turn the pump off and allow the wafer and silanizing agent to sit under vacuum overnight.! cautIon (Tridecafluoro-1,1,2,2-tetrahydrooctyl)-1-trichlorosilane is corrosive and toxic. Ensure that you wear proper PPE, and perform this inside a fume hood.

Fabrication of pDMs device ● tIMInG ~6 h16| Mount the SU-8 master in a 150-mm Petri dish by taping the edges of the wafer to the bottom of the Petri dish (Fig. 5).

17| Prepare two PDMS mixtures in 150-ml plastic cups and thoroughly mix them with a plastic spatula with the following specifications: device-layer PDMS, 15:1 prepolymer:cross-linker; support-layer PDMS 10:1 pre-polymer:cross-linker. It is suggested to make ~48 g of the device-layer PDMS mixture and ~66 g of the support-layer PDMS mixture.

18| Place both the device- and support-layer mixtures inside a vacuum desiccator, pump down the desiccation chamber with a vacuum pump for ~5 min and let it sit under vacuum for an additional 30 min. Degassing is required for removing bubbles that were trapped when mixing in Step 17. Once all bubbles are gone, remove the mixtures from the desiccator.

19| Pour the device-layer PDMS on top of the master so that it just covers the SU-8 features (e.g., ~1 mm deep of PDMS) (Fig. 5b). We recommend using compressed air to blow off any particulates that may have landed on the master before device-layer pouring. Particulates trapped within the PDMS cannot be removed after curing and can potentially interfere with image acquisition.

20| Degassing the PDMS-master. Degassing is necessary to remove bubbles that were trapped during PDMS pouring. Place the master inside the vacuum desiccator, pump down for ~5 min and let it sit under vacuum for an additional 15 min. crItIcal step During degassing, bubbles tend to rise and remain on top of the PDMS. To speed the degassing process up, we recommend using a compressed air source such as a duster to burst surface bubbles. This is done by gently blowing pulses of air at the surface of the wet PDMS (Fig. 5c). Once all bubbles are gone, remove the master from the desiccator. crItIcal step We recommend checking the device under a dissecting scope with top-side illumination to visually check for bubbles that may be trapped next to SU-8 features (Fig. 5d).

21| Place the master inside an oven at 70 °C for 20 min to cure the device layer. Remove the master from the oven. crItIcal step It is important that the device layer be cured enough so that it does not mix with the support layer in the next step. To check whether the device layer is cured, press the PDMS with your finger (in an area that is not over the device

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


features). If a ripple forms in the PDMS (Fig. 5e), place the master back in the oven for an additional 5 min.

22| Pour the support-layer PDMS on top of the cured device-layer PDMS for a PDMS device thickness of ~5 mm. Bubbles tend not to get trapped during the support-layer pouring; however, if it does occur, allow the master to sit while covered with a Petri dish for ~15 min. Bubbles should rise and can be taken care of with a duster as described in Step 19.

23| Place the master inside an oven at 70 °C for 4 h to cure the entire PDMS device. Remove it from the oven and allow it to cool for ~5 min.

24| Device peeling. By using a scalpel, cut the PDMS mold around the edge of the master wafer. Ensure that you cut all the way through the PDMS to the silicon wafer surface (Fig. 5f). By using the scalpel, wedge the blade into the cut and tilt it to peel a small section of PDMS mold from the master surface. Do this until enough PDMS is peeled to grip it with your hands. Peel the device by slowly pulling the PDMS mold up away from the master surface (Fig. 5g). Cut individual devices out with the scalpel.! cautIon Do not press the scalpel blade too hard into the master surface, because this may cause the master to fracture and risk complete loss of the master or may break the scalpel blade. Broken scalpel blades are a hazard as they tend to catapult into the air after fracture. Ensure that you wear eye protection. crItIcal step Do not peel the PDMS mold too quickly as this can cause delamination of the SU-8 structures or tearing of the PDMS mold (Fig. 4c).? trouBlesHootInG

25| Hole punching. Punch fluid inlet and outlet holes with a sharpened 19-gauge, blunt-tip needle. The needle must be sharpened (with a tool such as a Dremel) to punch a clean hole through the PDMS. Use the sharpened 19-gauge, blunt-tip needle to punch fluid inlet and outlet holes by pushing the needle all the way through the PDMS mold. Before retracting the needle back through the PDMS mold, make sure to remove the cylindrical PDMS cutout from the needle. Otherwise, during needle retraction the cutout will get lodged in the hole (Fig. 5h). Blow PDMS features with compressed air to remove particulates and follow that by covering the entire PDMS mold with Scotch Magic tape for protection until plasma bonding.? trouBlesHootInG

26| Plasma bonding. Place a clean 25 × 60 mm cover glass (i.e., sprayed with isopropyl alcohol and wiped with a Kimwipe, as well as blown with compressed air) inside the plasma chamber. Remove the tape from one of the PDMS molds and place the feature side up inside the plasma chamber (Fig. 5i). Place the plasma chamber cover back onto the plasma chamber with the valve closed; pump down the chamber to vacuum for 30 s. Turn on the plasma and wait until a purple glow is observed and stabilized (~5 s). Reduce the vacuum in the plasma chamber by opening the chamber cover valve slightly so that the purple glow becomes bright pink and make sure that the pink glow is stable. Let this sit for 20 s. Turn the plasma off, turn the pump off and release plasma chamber vacuum by completely opening the valve on the plasma chamber cover. Once atmospheric pressure is reached, the plasma chamber cover will fall off; you should be ready to catch it. Remove the cover glass from the plasma chamber with tweezers and place it on the clean bench top. Remove the PDMS mold from the

a b c

d e f

g h i

Figure 5 | Fabrication of the PDMS device. (a) SU-8 master fixed to the bottom of a 150-mm Petri dish with Scotch Magic tape. (b) Pouring PDMS onto the SU-8 master. (c) PDMS after degassing, with bubbles still present at the surface that are about to be removed by a duster. (d) Microscope image of bubbles trapped near the SU-8 features in device-layer PDMS. Scale bar, ~800 µm. (e) Testing the device layer after curing for 20 min. The ripple coming from the finger indicates that the device layer needs to be cured longer (~5 min). (f,g) Cutting the PDMS mold with a scalpel (f), which is followed by slowly peeling the PDMS mold from the master (g). (h) Hole punching. (i) PDMS mold and glass slide in the plasma chamber.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


plasma chamber by grasping at the sides of the device with tweezers and quickly place the feature side of the PDMS mold in direct contact with the cover glass for bonding. Gently press the PDMS mold into the cover glass, ensuring that a strong bond is formed. Allow it to sit for ~10 s. Check whether the plasma bonding was done correctly by trying to detach the PDMS mold from the cover glass. If the bonding was not done correctly, then the PDMS should be easily detached from the cover glass. Check bonding at all four corners of the device. crItIcal step Timing is important during plasma bonding. After plasma treatment, the PDMS mold needs to be attached to the glass slide in less than ~30 s. The activated PDMS surface will deactivate in air if not bonded in that time frame.? trouBlesHootInG

Device operation and imaging ● tIMInG ~0.5 h plus time for image acquisition27| Tubing connections: Use 1/32-inch i.d. × 3/32-inch o.d. silicone tubing for making the following connections (Fig. 2g–i). Connect the outlet of the pressure control system (i.e., the bare Luer needle on the stopcock valve) to the air inlet of the reservoir (i.e., the connection pin without the silicone tubing attached on the inside of the cap) (~60 cm). Connect the reservoir outlet to the microfluidic device (~60 cm). Make the connection between the silicone tubing and the microfluidic device with one of the connection pins (Fig. 2h). The pin should fit securely into the silicone tubing and the hole in the microfluidic device. Ensure that you place a manual pinch valve on the silicone tubing before making the connection. Connect the outlet of the microfluidic device to the waste collection bin. Make the connection between the silicone tubing and the microfluidic device with one of the connection pins (Fig. 2h). The pin should fit securely into the silicone tubing and the hole in the microfluidic device. Ensure that you place a manual pinch valve on the silicone tubing before making the connection.

28| Add ~40 ml of PBSB to the reservoir. Suspend the prepared embryos within the reservoir and close the cap tightly. See Box 1 for the immunostaining procedure we have commonly used.

29| Device degassing. The operation of the microfluidic device requires the microchannels to be entirely filled with the liquid mobile phase. Turn on the pressure source while making sure that the stopcock valve in the pressure control system is closed. Set the inlet pressure to ~15 p.s.i. by adjusting the regulator and monitoring the pressure gauge. Open the stopcock valve so that the fluid in the reservoir will flow into the device. Ensure that most of the air plugs are carried off the chip by convection (Fig. 6). This is most easily monitored under a dissecting scope. Gravity will cause the embryos to settle in the reservoir; they should settle far enough below the silicone tubing that they are not pulled onto the chip during degassing. In order to facilitate settling away from the silicone tubing, gently tap the sides of the reservoir. It should be easily seen that the embryo traps and resistance channels expand when increasing the device pressure from ambient to 15 p.s.i.

The deformation is crucial for embryo loading. Once it is noticed that most air plugs have been flown off the chip (~15 s after flow begins), close the a

c d

b

Figure 6 | Device loading. (a) Degassing the device before embryo loading. Most air plugs are carried off in the serpentine channel, but many air plugs remain within the cross-flow channels, which have to be removed via passive diffusion. Scale bar, ~1.5 mm. (b) Sequential frames showing a clogged group of embryos (red circles) being removed via probing with a 1-ml pipette tip. Scale bars, ~800 µm. The top left image highlights the targeted group, the bottom left image shows probing with a pipette tip, the top right image highlights the clogged group as it leaves in the serpentine channel, and the bottom right image shows the now-clear area where the clog was located. (c) Sequential frames tracking an individual embryo (red circles) as it approaches and orients into an embryo trap for end-on imaging. Scale bars, ~900 µm. Right, the oriented embryo in the final end-on position. (d) Image of the microfluidic device after loading with 71 embryos oriented for end-on imaging in the field of view. Scale bar, ~1 mm.

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


manual pinch valve at the microfluidic device outlet. Follow that by closing the manual pinch valve at the microfluidic device inlet. Turn off the pressure source by closing the stopcock valve. Allow the pressurized device to sit for ~15 min so that the remaining air plugs will be removed through passive diffusion through the PDMS walls. If the pinches adequately sealed the device, the embryo traps and resistance channels should remain expanded. crItIcal step It is crucial that all gas plugs be removed from the device, especially those located within the resistance channels. Air plugs within resistance channels will block the flow of the mobile phase and prevent embryos from correctly loading into embryo traps.? trouBlesHootInG

30| Device loading and imaging. Release the manual pinch valves at the microfluidic device inlet and outlet. Set the inlet pressure to ~6 p.s.i. and open the stopcock valve. Again, it should be easily seen that the embryo traps and resistance channels expand when increasing the device pressure from ambient to 6 p.s.i. The deformation is critical for embryo loading. Suspend the embryos in solution by quickly turning the reservoir upside down and back. Gravity will cause the embryos to settle toward the entrance of the silicone tubing so that they are carried onto the chip in a well-distributed manner. Embryos will sometimes stick to the PDMS walls. In order to release the stuck embryos, use a probe such as a 1-ml pipette tip to poke the PDMS device (Fig. 6b). Once all embryos are loaded onto the chip, turn off the inlet pressure by closing the stopcock valve and by turning off the pressure source (Fig. 6c,d). Replace the reservoir that previously held the embryos with another reservoir that contains 90% (vol/vol) glycerol solution. Flow the 90% glycerol solution into the microchannels by turning on the pressure source, setting the inlet pressure to ~6 p.s.i., opening the stopcock valve, allowing flow for ~1 min, closing the stopcock valve and turning off the pressure source. A concentration of 90% glycerol solution is used for imaging as it matches the refractive index of the glass slide. Disconnect the loaded device from the reservoir and waste collection and take it to the desired microscope for quantitative imaging of stained targets. Note that device loading and imaging do not have to be performed on separate microscopes. However, we suggest using a microscope that has a wide viewing area (e.g., dissecting scope) so that the entire device (or most of it) can be monitored during loading in order to detect loading errors such as clogging (Fig. 4d). crItIcal step Embryos should automatically orient and immobilize in the embryo traps for end-on imaging.? trouBlesHootInG

? trouBlesHootInGTroubleshooting advice can be found in table 1.

taBle 1 | Troubleshooting table.

step problem possible reason solution

4 Bubbles are present in SU-8 film

Wafer was too hot when dispensing SU-8

It is important to allow the wafer ample time (~5 min) to cool after being exposed to a hot plate. Rule of thumb: cool to the touch

Bubbles were trapped when pouring SU-8 onto wafer

Pour the SU-8 as close to the wafer surface as possible and keep the stream in one spot at the center; do not attempt to cover the entire wafer when pouring

6 SU-8 film appears cracked after exposure

The film was overexposed Decrease the energy dose by ~10%

8 Bubbles are present in SU-8 film

Wafer was too hot when dispensing SU-8

It is important to allow the wafer ample time (~5 min) to cool after being exposed to a hot plate. Rule of thumb: cool to the touch

Bubbles were trapped when pouring SU-8 onto wafer

Pour the SU-8 as close to the wafer surface as possible and keep the stream in one spot at the center; do not attempt to cover the entire wafer when pouring

(continued)

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


taBle 1 | Troubleshooting table (continued).


11 Features are not visible after exposure

The film was underexposed Increase the energy dose by ~10%

12 Features appear after postexpose bake but are distorted

Missing adhesion layer It is recommended to use an adhesion layer as described in Steps 4–7

13 Features are missing (Fig. 4b)

The film was overdeveloped Develop for ~10% less time. It is suggested to check the progress of development with isopropyl alcohol frequently

The film was underexposed Increase the energy dose by 10%

Mask has some pattern errors It is recommended to inspect the quality of the mask after purchase

Missing adhesion layer It is recommended to use an adhesion layer as described in Steps 4–7

24 PDMS mold tears PDMS mold is undercured Be sure to cure in Step 23 for 4 h. If done so, cure for an additional 30 min

PDMS mold cannot be cleanly pealed from master surface (Fig. 4c)

PDMS mold is undercured (if this is the problem, usually small areas of PDMS mold cannot be cleanly pealed)

Be sure to cure in Step 23 for 4 h. If necessary, cure for an additional 30 min

Master was not adequately silanized (if this is the problem, usually large areas of PDMS mold cannot be cleanly pealed)

Be sure to silanize the master as described in Step 15. Silanizing agent might have deactivated (deactivates in the present of oxygen) and will require new reagent

25 PDMS mold tears during hole punching

19-gauge blunt tip needle is not sharp enough

Be sure to sharpen the 19-gauge blunt tip needle with a Dremel tool

Inlet/outlet hole appears narrow

19-gauge blunt tip needle was damaged during sharpening

Usually when sharpening, the needle tip gets pushed inward, thus narrowing the inner diameter. Insert the tip of the surgical blade into the needle tip and twist to regain the original inner diameter of the needle

26 PDMS mold does not bond to the cover glass

Cover glass was not clean Be sure to clean the cover glass with isopropyl alcohol and remove particulates with a duster

PDMS mold was not clean Use clear adhesive tape to remove particulates from the PDMS surface

Plasma treatment did not success-fully activate the PDMS surface

Perform plasma treatment for longer time (~5 s more)

PDMS surface was deactivated before contact between PDMS mold and cover glass was made

Make sure once plasma treatment is completed that the cover glass and PDMS mold are brought into contact for bonding in less than 30 s

29 Traps do not appear to deform under pressure

PDMS is too stiff Make sure the device layer is sufficiently cured that it does not mix with the support layer when the support layer is poured. The device layer should not appear to flow when pouring support layer (undercured). If this is noticed, bake for longer time (~5 min more)

(continued)

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


● tIMInGSteps 1–15, microfabrication of SU-8 master: ~5 h plus overnight silanizationSteps 16–26, fabrication of PDMS device: ~6 hSteps 27–30, device operation and imaging: ~0.5 h plus time for image acquisitionBox 1, embryo immunostaining: ~7 h plus overnight incubation

antIcIpateD resultsThe microfluidic device developed from this protocol should be able to orient and immobilize embryos for end-on imaging in a massively parallel manner (Fig. 6d). The fabrication of the SU-8 master can be made by a regular clean-room user after one full run-through of PROCEDURE Steps 1–15. Although the fabrication scheme is not error proof, especially from the multiple coats of SU-8 that are required to attain a film thickness of 500 µm, a functional device can still be easily produced even with occasional minor errors. For example, a device can successfully load a large volume of embryos even with nonuniform film thickness across the SU-8 master. Practice and patience usually result in better yield in the microfabrication steps. The limiting factor to whether a device is successful is primarily dependent on how well the master is fabricated. To attain

taBle 1 | Troubleshooting table (continued).


30 Embryos are clogging in the tubing before reaching the microfluidic device

Wrong size tubing was used Make sure to use silicone tubing (1/32-inch i.d. × 3/32-inch o.d.)

The connection pins were accidently pinched during equipment setup

Do not grip the hollow steel pins too tightly when separating the plastic Luer end from the needle

Inlet/outlet holes were not successfully punched

Make sure to reopen the needle tip after sharpening, or the needle will cut a hole that is narrower than required

Embryo suspension density is too high

Suspend embryos from one collection from one population (~700 embryos) in at least 40 ml of mobile phase (or even up to 100 ml)

Embryos are clogging within the serpentine channel (Fig. 4d)

No surfactant used It is recommended to use a biocompatible surfactant (e.g., 1% (wt/vol) BSA) to prevent embryos from sticking to each other and device walls

Embryo suspension is too high a density

Suspend embryos from one collection from one population (~700 embryos) in at least 40 ml of mobile phase or even up to 100 ml

Embryos do not get trapped

PDMS is too stiff Make sure device layer is sufficiently cured that it does not mix with the support layer when the support layer is poured. The device layer should not appear to flow when pouring support layer (undercured). If this is noticed, bake for longer time (~5 min more)

Device was not properly degassed and air plugs are clogging cross-flow channels

Make sure to degas for 15 min at ~15 p.s.i. To check if air bubbles are still present one can vary the inlet pressure or open/close outlet pinch valve. This should cause plugs (if present) to move slightly, which can be seen. If plugs are noticed, degas for longer time (~15 min more)

No fluorescence is detected

Target did not successfully get labeled with fluorescent marker

Make sure to follow the immunostaining procedure outlined in Box 1 (or whatever technique was used to label embryos)

©20

13 N

atu

re A

mer

ica,

Inc.

All

rig

hts

res

erve

d.

protocol


high loading efficiencies (i.e., > 90%), one must be particularly careful with the fidelity of the SU-8 structures. However, this is often not a requirement as our device has > 700 traps and very modest loading efficiencies (e.g., 20%) can still yield ~150 end-on–oriented embryos. Even with a 5% loading efficiency, one would have a vast improvement with regard to sample size over previous studies that have observed ~10 embryos. Typically, devices that have mild defects will still load consistently at ~20% efficiency. Thus, a novice user can make a functioning master in the first run-through of this protocol.

In general, this microfluidic device is unique with the ability to immobilize and orient oblong embryos within its broad applications. As the embryos are physically immobilized within the embryo traps, the device can be transported without loss of the end-on orientation of embryos. This is particularly useful when loading the device on a dissecting microscope at low magnification and transporting the loaded device to more powerful microscopy setups elsewhere, such as confocal microscopes. The transportability is also useful in that loading can be done by one individual (e.g., microfluidics ‘expert’) and taken to another for high-quality imaging (e.g., imaging ‘expert’). This device allows for the immunostaining of embryos in order to visualize many morphogens, including Dorsal as we have done previously23. In the future, we also envision that many other biochemical techniques, which have already been adapted to Drosophila development (e.g., FISH9), will be used in preparing embryos and imaging on the microfluidic device.

Note: Supplementary information is available in the online version of the paper.

acknowleDGMents This work was supported by the US National Science Foundation (DBI-0649833 to H.L., EFRI 1136913 to S.Y.S. and H.L.) and the US National Institutes of Health (grant nos. NS058465 to H.L. and GM078079 to S.Y.S.). H.L. is a DuPont Young Professor and a Sloan Research Fellow. We thank J. Ding for technical assistance.

autHor contrIButIons T.J.L. and M.Z. fabricated the devices and tested the protocol. H.L. and S.Y.S. oversaw the project. T.J.L. wrote the paper. All authors (T.J.L., M.Z.,B.L., S.Y.S. and H.L.) contributed to the design of the experiments and edited the manuscript.

coMpetInG FInancIal InterestsThe authors declare no competing financial interests.

Reprints and permissions information is available online at http://www.nature.com/reprints/index.html.

1. Bier, E. Drosophila, the golden bug, emerges as a tool for human genetics. Nat. Rev. Genet. 6, 9–23 (2005).

2. Adams, M.D. et al. The genome sequence of Drosophila melanogaster. Science 287, 2185–2195 (2000).

3. Gurdon, J.B. & Bourillot, P.Y. Morphogen gradient interpretation. Nature 413, 797–803 (2001).

4. Frohnhofer, H.G. & Nusslein-Volhard, C. Organization of anterior pattern in the Drosophila embryo by the maternal gene bicoid. Nature 324, 120–125 (1986).

5. Roth, S., Stein, D. & Nusslein-Volhard, C. A gradient of nuclear localization of the dorsal protein determines dorsoventral pattern in the Drosophila embryo. Cell 59, 1189–1202 (1989).

6. Steward, R. Relocalization of the Dorsal protein from the cytoplasm to the nucleus correlates with its function. Cell 59, 1179–1188 (1989).

7. Rushlow, C.A. & Shvartsman, S.Y. Temporal dynamics, spatial range, and transcriptional interpretation of the Dorsal morphogen gradient. Curr. Opin. Genet. Dev. 22, 542–546 (2012).

8. Chung, K. et al. A microfluidic array for large-scale ordering and orientation of embryos. Nat. Methods 8, 171–176 (2011).

9. Kanodia, J.S. et al. A computational statistics approach for estimating the spatial range of morphogen gradients. Development 138, 4867–4874 (2011).

10. Kanodia, J.S. et al. Pattern formation by graded and uniform signals in the early Drosophila embryo. Biophys. J. 102, 427–433 (2012).

11. Helman, A. et al. RTK signaling modulates the Dorsal gradient. Development 139, 3032–3039 (2012).

12. Kim, Y. et al. Gene regulation by MAPK substrate competition. Dev. Cell 20, 880–887 (2011).

13. Crane, M.M., Chung, K. & Lu, H. Computer-enhanced high-throughput genetic screens of C. elegans in a microfluidic system. Lab Chip 9, 38–40 (2009).

14. Chung, K., Crane, M.M. & Lu, H. Automated on-chip rapid microscopy, phenotyping and sorting of C. elegans. Nat. Methods 5, 637–643 (2008).

15. Crane, M.M., Chung, K., Stirman, J. & Lu, H. Microfluidics-enabled phenotyping, imaging, and screening of multicellular organisms. Lab Chip 10, 1509–1517 (2010).

16. Liang, H.L. et al. The zinc-finger protein Zelda is a key activator of the early zygotic genome in Drosophila. Nature 456, 400–403 (2008).

17. Li, X.Y. et al. Transcription factors bind thousands of active and inactive regions in the Drosophila blastoderm. PLoS Biol. 6, e27 (2008).

18. Reeves, G.T. et al. Dorsal-ventral gene expression in the Drosophila embryo reflects the dynamics and precision of the dorsal nuclear gradient. Dev. Cell 22, 544–557 (2012).

19. Chung, K.H., Rivet, C.A., Kemp, M.L. & Lu, H. Imaging single-cell signaling dynamics with a deterministic high-density single-cell trap array. Anal. Chem. 83, 7044–7052 (2011).

20. Akagi, J. et al. Miniaturized embryo array for automated trapping, immobilization and microperfusion of zebrafish embryos. PLoS ONE 7, 36630 (2012).

21. Kohli, V. et al. An alternative method for delivering exogenous material into developing zebrafish embryos. Biotechnol. Bioeng. 98, 1230–1241 (2007).

22. Samara, C. et al. Large-scale in vivo femtosecond laser neurosurgery screen reveals small-molecule enhancer of regeneration. Proc. Natl Acad. Sci. USA 107, 18342–18347 (2010).

23. Tomer, R., Khairy, K., Amat, F. & Keller, P.J. Quantitative high-speed imaging of entire developing embryos with simultaneous multiview light-sheet microscopy. Nat. Methods 9, 755–U181 (2012).

24. Keller, P.J., Schmidt, A.D., Wittbrodt, J. & Stelzer, E.H.K. Reconstruction of zebrafish early embryonic development by scanned light sheet microscopy. Science 322, 1065–1069 (2008).

25. Fakhoury, J.R., Sisson, J.C. & Zhang, X. Microsystems for controlled genetic perturbation of live Drosophila embryos: RNA interference, development robustness and drug screening. Microfluid. Nanofluid. 6, 299–313 (2009).

26. Haywood, A.F. & Staveley, B.E. Parkin counteracts symptoms in a Drosophila model of Parkinson’s disease. BMC Neurosci. 5, 14 (2004).

27. Liberman, L.M., Reeves, G.T. & Stathopoulos, A. Quantitative imaging of the Dorsal nuclear gradient reveals limitations to threshold-dependent patterning in Drosophila. Proc. Natl Acad. Sci. USA 106, 22317–22322 (2009).

28. Kanodia, J.S. et al. Dynamics of the Dorsal morphogen gradient. Proc. Natl Acad. Sci. USA 106, 21707–21712 (2009).

29. Witzberger, M.M., Fitzpatrick, J.A., Crowley, J.C. & Minden, J.S. End-on imaging: a new perspective on dorsoventral development in Drosophila embryos. Dev. Dyn. 237, 3252–3259 (2008).

30. Markow, T.A., Beall, S. & Matzkin, L.M. Egg size, embryonic development time and ovoviviparity in Drosophila species. J. Evolution. Biol. 22, 430–434 (2009).

31. Aboobaker, A.A., Tomancak, P., Patel, N., Rubin, G.M. & Lai, E.C. Drosophila microRNAs exhibit diverse spatial expression patterns during embryonic development. Proc. Natl Acad. Sci. USA 102, 18017–18022 (2005).

32. Ashburner, M. Grape-apple juice agar plates. In Drosophila Protocols (eds. Ashburner, M., Hawley, R.S. & Sullivan, W.) 658 (Cold Spring Harbor Laboratory Press, 2000).

http://www.nature.com/doifinder/10.1038/nprot.2013.034

http://www.nature.com/reprints/index.html

http://www.nature.com/reprints/index.html

Microfluidic trap array for massively parallel imaging of...

Documents

Transcript of Microfluidic trap array for massively parallel imaging of...