We thank Dr. Krishanu Ray for Drosophila stocks, Tarjani Agarwal for maintaining a Drosophila cage, Peter Juo for nuIs25 and CGC for C. elegans strains. SPK made jsIs609 in Michael Nonet's laboratory. We thank Arpan Agnihotri (BITS Pilani) for his help in time-lapse imaging of mitochondria transport of jsIs609 animals in microfluidic devices. We are grateful to Dr. Vatsala Thirumalai and Surya Prakash for providing us with zebrafish embryos. We thank Dr. Krishna and CIFF at NCBS for use of the spinning disc confocal microscope supported by the Department of Science and Technology- Centre for Nanotechnology (No. SR/55/NM-36-2005). We also thank Kaustubh Rau, V. Venkatraman and Chetana Sachidanand for discussions. This work was funded by the DBT post-doctoral fellowship (S.M.), DST fast-track scheme (S.M.) and a DBT grant to (S.P.K.). S.A. was supported by DST and CSIR grants to SPK.

1. SU8 Master Fabrication
<ol>
 <li>Design the microfluidic structures using Clewin software and print it using 65,024 DPI laser plotter with minimum feature size of 8 &mu;m on circuit board film. </li>
 <li>Clean 2 cm X 2 cm silicon wafers with native oxide in 20% KOH for 1 min and rinse in deionized water; one wafer each for the flow layer and its corresponding control layer. </li>
 <li>Blow dry the pieces with nitrogen gas and dehydrate on a hot plate at 120 &deg;C for 4 hr. Allow the pieces to cool down to room temperature before proceeding to the next step.</li>
 <li>Place silicon pieces one at a time on the spinner chuck and turn on the vacuum. Cover the silicon surfaces with ~20 &mu;l hexamethyl-disilazane (HMDS) and coat them using a SPIN150 spinner at 500 rpm for 5 sec followed by 3,000 rpm for 30 sec.</li>
 <li>Cover silicon wafers completely with ~1.5 ml of SU8-2025 (<a href="http://www.microchem.com/Prod-SU82000.htm" target="_blank">http://www.microchem.com/Prod-SU82000.htm</a>) and coat wafers using SPIN150 spinner at 500 rpm for 5 sec followed by 2,000 rpm for 30 sec. This spinning protocol produces a photoresist thickness of ~40 &mu;m that is suitable for the control and flow layers for early larval stages of C. elegans.</li>
 <li>Spin ~1.5 ml SU8-2050 using SPIN150 spinner at 500 rpm for 5 sec followed by 2,000 rpm for 30 sec to obtain a photoresist thickness of ~80 &mu;m that is suitable for flow layer fabrication for late larval stages of C. elegans, basal flow layer for Drosophila/zebrafish larvae and their corresponding control layers.</li>
 <li>Place the silicon pieces on hot plate with the SU8 coated layer on top. Soft bake the coated silicon pieces at 65 &deg;C for 1 min followed by 95 &deg;C for 10 min. Allow the pieces to cool down to room temperature before proceeding to the next step. </li>
 <li>Place the soft-baked silicon pieces on the exposure stage with SU8-2025 coated surface on the top facing the UV lamp. Use the photo mask with design I (L1, Figure 1B) and design II (L2, Figure 1B) for the flow layer and control layer pattern respectively for early larval stages of C. elegans. Place the photo mask on top of the SU8 layer and ensure the mask is flat against the coated layer. Open the shutter of the UV lamp and expose the soft-baked SU8 wafers to 200 Watt UV lamp through the photo mask for 15 sec.</li>
 <li>Use the photo mask with design I (L1, Figure 1B) and design II (L2, Figure 1B) on SU8-2050 coated pieces (prepared in step 1.6) to fabricate the flow layer and control layer for late larval stages of C. elegans. Use the photo mask with the design I (L1, Figure 1C) and design II (L2, Figure 1C) for the basal flow layer and control layer respectively for Drosophila/zebrafish larvae on wafers coated with SU8-2050 (prepared in step 1.6). Expose the SU8 surfaces through the photo mask to the 200 Watt UV lamp for 15 sec.</li>
 <li>Place the exposed silicon pieces on a hot plate with the coated layer on top. Post bake the wafers at 65 &deg;C for 1 min followed by 95 &deg;C for 10 min. Allow the pieces to cool down before proceeding to the next step. </li>
 <li>Develop the pieces using the SU8 developer solution for 20 min. Rinse the pieces in Iso-Propyl Alcohol (IPA) and blow dry using nitrogen gas.</li>
 <li>Place the silicon pieces in a desiccator with the SU8 pattern on top. Coat the pieces with 50 &mu;l of tricholoro (1H,1H,2H,2H-perfluorooctyl) silane vapor in a desiccator for 2 hr.</li>
</ol>
Precautions: Take safety precautions for chemical handling during KOH treatment, photoresist coating and developing. Silane vapor coating should be performed inside a desiccator in a ventilated area. Protect SU8 resist from over exposure to light. Use protective eyewear with an UV light source.
2. PDMS Mold Fabrication
<ol>
 <li>Prepare 10:1 PDMS by combining 50 g of Sylgard 184 base with 5 g of curing agent in a plastic cup. Mix the content manually for ~3 min. Degas the mixture inside a desiccator to remove all air bubbles formed during mixing.</li>
 <li>Pour a 5 mm thick PDMS mixture over the silicon wafers with SU8 pattern of design II, for control layer, gently to avoid formation of air bubbles. In case bubbles are formed during pouring, degas the PDMS on the SU8 pattern in low vacuum to remove all the bubbles.</li>
 <li>Place the silicon wafers with SU8-2025 pattern of design I, for the flow layer, on spinner chuck and turn on the vacuum to hold them. Cover the wafers with ~1 ml of PDMS mix and coat the wafers using SPIN150 spinner at 500 rpm for 5 sec followed by 1,500 rpm for 30 sec.</li>
 <li>Coat ~1 ml PDMS mix on silicon wafers with the 80 &mu;m SU8-2050 pattern of design I (L1, Figure 1B), for the flow layer for late larval stages of C. elegans using the SPIN150 spinner at 500 rpm for 5 sec followed by 1,000 rpm for 30 sec. Coat a thick layer of PDMS mix on the silicon pieces with SU8-2050 pattern of flow layer design I (L1, Figure 1C) for Drosophila/zebrafish larvae, using SPIN150 spinner at 500 rpm for 35 sec.</li>
 <li>Bake PDMS coated silicon pieces of design I and wafers with corresponding design II for control layer containing PDMS for C. elegans and/or Drosophila/zebrafish larvae in a hot air convection oven at 50 &deg;C for 6 hr.</li>
 <li>Cut out the PDMS piece from the silicon substrate containing the SU8 pattern II for C. elegans and/or Drosophila/zebrafish larvae using a sharp blade. Punch a small hole of ~1 mm diameter on top of the reservoir connecting the main trap in the PDMS mold using a Harris puncher.</li>
 <li>Place the punched PDMS block (containing the 40 &mu;m thick mold for control layer L2) on a plastic tray, with the molded side of the control layer facing up. Place the PDMS coated silicon wafer containing the 40 &mu;m thick SU8 design I on the same tray, with the PDMS coated surface facing up. Insert the tray inside the chamber of plasma cleaner and turn on the vacuum for 2 min. Subsequently, turn on the plasma power and lower the chamber pressure until the chamber turns light pink in color. Expose both surfaces to 18 Watt air plasma under low vacuum for 2 min.</li>
 <li>Place the punched PDMS block removed from the 80 &mu;m thick SU8 master containing pattern II for late larval stages of C. elegans or Drosophila/zebrafish larvae on a plastic tray, with the molded side of the control layer facing up. Place the corresponding baked PDMS layer spin coated on the silicon wafer containing 80 &mu;m thick design I for later C. elegans stages or Drosophila/zebrafish larvae simultaneously on the same tray with the flat PDMS coated surface facing up. Use the same protocol mentioned above to expose them to air plasma.</li>
 <li>Place the two plasma treated surfaces for C. elegans and/or Drosophila/zebrafish larvae together with gentle pressure and bake them in hot air convection oven at 50 &deg;C for 2 hr.</li>
 <li>Cut out the bonded devices from the silicon substrate with SU8 design I for C. elegans and/or Drosophila/zebrafish larvae. Punch access holes at the inlet and outlet reservoirs of the flow channel.</li>
 <li>Place the bonded PDMS mold on a plastic tray for C. elegans, with the molded side of the flow layer design facing up. Clean glass cover slip (22 X 22 mm, No. 1 thickness) and place it on the same plastic tray. Insert the tray containing PDMS mold and glass cover slip inside the plasma chamber. Expose the bottom PDMS surface of the device and glass cover slip to 18 Watt air plasma at low pressure for 2 min. Place the two plasma treated surfaces with gentle pressure and bake them in hot air convection oven at 50 &deg;C for 2 hr.</li>
 <li>Store the device in a clean environment for future use. </li>
</ol>
3. Additional Steps for Drosophila/Zebrafish Device
<ol>
 <li>Expose clean glass surface of size 2 cm X 2 cm with 50 &mu;l of tricholoro(1H,1H,2H,2H-perfluorooctyl) silane vapor in a desiccator for 2 hr.</li>
 <li>Spin ~1 ml of 10:1 PDMS mix prepared in step 2.1 on the silanized glass surface using SPIN150 spinner at 500 rpm for 35 sec. Bake the PDMS coated glass substrates in a hot air oven at 50 &deg;C for 6 hr with the coated surface facing up. Allow the PDMS layer to cool down to room temperature before proceeding to the next step.</li>
 <li>Punch the coated layer using a custom built sharp metal puncher at the middle of the PDMS, to form the PDMS spacer layer. The shape of the metal puncher is designed similar to the flow design for Drosophila/zebrafish (L1, Figure 1C).</li>
 <li>Expose the PDMS spacer layer and the single-bonded block (flow layer and control layer, made in step 2.9 for Drosophila/zebrafish larvae) to 18 Watt air plasma using plasma cleaner at low vacuum. Place the two plasma treated surfaces together and bake them in a hot air oven at 50 &deg;C for 2 hr. </li>
 <li>Cut the bonded device from the glass, punch access holes at the inlet and outlet reservoirs and bond it to a glass cover slip (22 X 22 mm, No. 1 thickness) using a plasma cleaner as mentioned in step 2.11. This would produce an immobilization device for first instar Drosophila larvae with a channel height of ~500 &mu;m. For zebrafish larvae, duplicate the PDMS spacer layer fabrication process with an additional bonding step. The dual layer of PDMS-S produces a channel height of 900 &mu;m for 30 hfp zebrafish larvae. </li>
</ol>
Precautions: Avoid dust particles during device fabrication. Two plasma cleaned surfaces need to be completely dust free for proper bonding. Store the devices in a desiccator in situations where a special clean room is not available in order to minimize accumulation of dust particles in devices.
4. Using PDMS Membrane
<ol>
 <li>Connect one end of a micro-flex tube (inner diameter ~1.6 mm, outer diameter ~4.8 mm) to a pressurized nitrogen gas supply regulator and the other end to the main trap reservoir through an 18 gauge needle (outer diameter ~1.25 mm) glued to the tube. A 3-way stopcock used in the middle of the tube connection allows application or release of pressure on the membrane.</li>
 <li>Fill the tube connected to the PDMS device with a 10 cm column of distilled water before connecting it to the reservoir of the main trap of a C. elegans and/or Drosophila/zebrafish larvae device. </li>
 <li>Turn the valve of the nitrogen supply to read 14 psi and monitor the membrane of the main trap deflecting towards the flow channel at low magnification of an inverted microscope. Length of the water column in the micro-flex tube is compressed whenever the 3-way stopcock is open. Edges of the deflected membrane are visible in transmitted light (Figure 1I and 1J). Membrane deflection causes displacement of dust particles/bubbles under the PDMS membrane and pushes them to the channel boundaries.</li>
 <li>Wait till the liquid front fills the microchannel completely without any trapped air. </li>
 <li>Release the pressure by using the 3-way stopcock to relax the membrane to its resting position. </li>
</ol>
5. Inserting C. elegans, Drosophila and Zebrafish Larvae into the Device and Immobilizing Them Under the Flexible PDMS Membrane 
<ol>
 <li>Fill the flow channel with M9 buffer [3 g KH2PO4, 6 g Na2HPO4, 5 g NaCl, 1 ml 1 M MgSO4, H2O to 1 L, sterilize by autoclaving] for C. elegans 18 or 1X PBS [8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2PO4, H2O to 1 L, sterilize by autoclaving] for Drosophila/zebrafish larvae 19 for 10 min before an experiment.</li>
 <li>Locate a single C. elegans of required stage on a NGM plate 18 (or first instar Drosophila larvae from agar egg plate or manually dechorionated 30 hpf zebrafish larvae in liquid medium 20) using a low magnification stereo microscope. Pick a single animal using a small volume of buffer solution into a micro tip. Use a micro tip with its end cut to accommodate the larger Drosophila or zebrafish larvae in buffer. </li>
 <li>Push the single organism through the flow channel inlet. Adjust the pipetting pressure to position the individual animal under the main trap. </li>
 <li>Increase the pressure of the PDMS membrane slowly to position the animal against the channel boundary and immobilize it with 14 psi pressure for C. elegans, 7 psi for Drosophila and 3 psi for zebrafish larvae.</li>
 <li>Position the immobilized animal at the center of the microscopic field of view. Use an inverted microscope at desired settings for high resolution bright field or fluorescence imaging. Acquire single or time-lapse fluorescence images at predefined frame rate.</li>
 <li>Release the trap pressure and monitor the locomotion of the animal for 5-10 min at low magnification. </li>
 <li>Flush the animal and insert a fresh animal to repeat the above steps. </li>
 <li>Wash all the animals from the waste reservoir using distilled water applied using a syringe. Dry the channel by pushing air using a syringe for future reuse. </li>
</ol>
Precautions: Animals requiring higher pipetting pressure to flow inside the main channel tend to show poor locomotion/health after release from the trap and should be avoided for imaging. Clean the flow channel for any trace of buffer to prevent clogging of channel by crystals. If oil is used for high resolution imaging, clean the glass cover slip to be able to maintain good signal to noise ratio for subsequent experiments.

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+regulation+of+bidirectional+mitochondrial+transport+is+coordinated+with+axonal+outgrowth.">The regulation of bidirectional mitochondrial transport is coordinated with axonal outgrowth.</a> J. Cell. Sci. 104, 917-927 (1993).</li><li>Louie, K., Russo, G. J., Salkoff, D. B., Wellington, A., Zinsmaier, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+imaging+conditions+on+mitochondrial+transport+and+length+in+larval+motor+axons+of+Drosophila.">Effects of imaging conditions on mitochondrial transport and length in larval motor axons of Drosophila.</a> Comp. Biochem. Physiol. A. Mol. Integr. Physiol. 151, 159-172 (2008).</li><li>Pilling, A. D., Horiuchi, D., Lively, C. M., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin-1+and+Dynein+are+the+primary+motors+for+fast+transport+of+mitochondria+in+Drosophila+motor+axons.">Kinesin-1 and Dynein are the primary motors for fast transport of mitochondria in Drosophila motor axons.</a> Mol. Biol. Cell. 17, 2057-2068 (2006).</li></ol>

No conflicts of interest declared.

PDMS microfluidic devices are optically transparent therefore can be used for high resolution in vivo imaging of any transparent/translucent model organism. Our design is suitable for high magnification spatio-temporal imaging of cellular and sub-cellular events in intact live animals. Microfabrication using soft lithography techniques allows easy manipulation of device dimensions for various sizes of model organisms. Devices of various sizes are fabricated for different stages of C. elegans, Drosop...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Silicon wafers</td>
 <td>University wafer</td>
 <td>150 mm (100) Mech Grade SSP Si</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Clewin Software</td>
 <td>WieWeb software</td>
 <td>Version 2.90</td>
 <td>&nbsp;</td>
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 <tr>
 <td>Laser plotter</td>
 <td>Fine Line Imaging</td>
 <td>65,024 DPI</td>
 <td>&nbsp;</td>
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 <tr>
 <td>HMDS</td>
 <td>Sigma-Aldrich</td>
 <td>440191-100ML</td>
 <td>&nbsp;</td>
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 <tr>
 <td>SU8</td>
 <td>Microchem</td>
 <td>SU8-2025, SU8-2050</td>
 <td>&nbsp;</td>
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 <td>Developer</td>
 <td>Microchem</td>
 <td>SU8 Developer</td>
 <td>&nbsp;</td>
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 <td>Silane</td>
 <td>Sigma-Aldrich</td>
 <td>448931-10G</td>
 <td>&nbsp;</td>
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 <td>PDMS</td>
 <td>Dow corning</td>
 <td>Sylgard 184</td>
 <td>&nbsp;</td>
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 <td>UV lamp</td>
 <td>Oriel</td>
 <td>66943</td>
 <td>200W Hg Oriel Light</td>
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 <td>Hot air oven</td>
 <td>Ultra Instruments</td>
 <td>Custom made</td>
 <td>Set at 50 &deg;C</td>
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 <td>Hot plate</td>
 <td>IKA Laboratory Equipment</td>
 <td>3810000</td>
 <td><a href="http://www.ika.com" target="_blank">http://www.ika.com</a></td>
 </tr>
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 <td>Plasma cleaner</td>
 <td>Harrick Plasma</td>
 <td>PDC-32G</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Spinner</td>
 <td>Semiconductor Production Systems</td>
 <td>SPIN150-NPP</td>
 <td><a href="http://www.SPS-Europe.com" target="_blank">www.SPS-Europe.com</a></td>
 </tr>
 <tr>
 <td>Glass cover slip</td>
 <td>Gold Seal</td>
 <td>22 X 22 mm, No. 1 thickness</td>
 <td>&nbsp;</td>
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 <tr>
 <td>C. elegans</td>
 <td>Caenorhabditis Genetics Center (CGC)</td>
 <td>e1265, ayIs4</td>
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 <td>Drosophila</td>
 <td>Bloomington</td>
 <td>P{chaGAL4}/cyo, UAS-syt.eGFP</td>
 <td>&nbsp;</td>
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 <td>Zebrafish</td>
 <td>Indian wild type</td>
 <td>Wild type</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Tygon tube</td>
 <td>Sigma</td>
 <td>Z279803</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micro needle</td>
 <td>Sigma</td>
 <td>Z118044</td>
 <td>Cut into 1 cm pieces</td>
 </tr>
 <tr>
 <td>3-way stopcock</td>
 <td>Sigma</td>
 <td>S7521</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Harris puncher</td>
 <td>Sigma</td>
 <td>Z708631</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Compressed nitrogen gas</td>
 <td>Local Gas supplier</td>
 <td>&nbsp;</td>
 <td>Use a regulator to control the pressure</td>
 </tr>
 <tr>
 <td>Stereo microscope</td>
 <td>Nikon</td>
 <td>SMZ645</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Confocal microscope</td>
 <td>Andor &amp; Olympus</td>
 <td>Yokogawa spinning disc confocal microscope</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>ImageJ</td>
 <td>National Institutes of Health</td>
 <td><a href="http://www.rsbweb.nih.gov/ij" target="_blank">www.rsbweb.nih.gov/ij</a> </td>
 <td>Java based image processing program</td>
 </tr>
 </tbody>
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simple microfluidic devices for in vivo imaging of c. elegans, drosophila and zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

The immobilization device is a bilayer PDMS block fabricated by bonding two layers: a flow layer (Layer 1) and a control layer (Layer 2) as shown in Figure 1. The main trap is connected to a nitrogen gas cylinder through a regulator and a 3-way stop cock to apply necessary (3-14 psi) pressure onto the membrane through a liquid column (Figure 1A). The deflected membrane immobilizes C. elegans, Drosophila or zebrafish larvae in the flow channel designed with different dim...

Watch this Scientific Journal Video about Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish at JoVE.com

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

simple-microfluidic-devices-for-vivo-imaging-c-elegans-drosophila

Research

JoVE Journal

Bioengineering

17.0K Views.  NCBS-TIFR. Un semplice dispositivo microfluidico è stato sviluppato per eseguire anestetico libera In vivo Imaging C. elegans, Intatto Drosophila Larve e larve di zebrafish. Il dispositivo utilizza una membrana deformabile PDMS per immobilizzare questi organismi modello per eseguire lapse imaging tempo di numerosi processi come il battito cardiaco, la divisione cellulare e subcellulare trasporto neuronale. Abbiamo illustrato l'utiliz...

Video: Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Separating Beads and Cells in Multi-channel Microfluidic Devices Using Dielectrophoresis and Laminar Flow

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting and counting cells. However, manipulating the cell within the fluidic domain remains a challenge and requires complicated fabrication protocols for forming valves and electrodes, or demands specialty equipment like optical tweezers. Here, we demonstrate that conventional printed circuit boards (PCB) can be used for the non-contact manipulation of cells by employing dielectrophoresis (DEP) for bead and cell manipulation in laminar flow fields for bioactuation, and for cell and bead separation in multichannel microfluidic devices. First, we present the protocol for assembling the DEP electrodes and microfluidic devices, and preparing the cells for DEP. Then, we characterize the DEP operation with polystyrene beads. Lastly, we show representative results of bead and cell separation in a multichannel microfluidic device. In summary, DEP is an effective method for manipulating particles (beads or cells) within microfluidic devices.

Dielectrophoresis (DEP) is an effective method to manipulate cells. Printed circuit boards (PCB) can provide inexpensive, reusable and effective electrodes for contact-free cell manipulation within microfluidic devices. By combining PDMS-based microfluidic channels with coverslips on PCBs, we demonstrate bead and cell manipulation and separation within multichannel microfluidic devices.

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting ...

A Microfluidic Technique to Probe Cell Deformability

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an efficient manner. Typically, data for ~102 cells can be acquired within a 1 hr experiment. An automated image analysis program enables efficient post-experiment analysis of image data, enabling processing to be complete within a few hours. Our device geometry is unique in that cells must deform through a series of micron-scale constrictions, thereby enabling the initial deformation and time-dependent relaxation of individual cells to be assayed. The applicability of this method to human promyelocytic leukemia (HL-60) cells is demonstrated. Driving cells to deform through micron-scale constrictions using pressure-driven flow, we observe that human promyelocytic (HL-60) cells momentarily occlude the first constriction for a median time of 9.3 msec before passaging more quickly through the subsequent constrictions with a median transit time of 4.0 msec per constriction. By contrast, all-trans retinoic acid-treated (neutrophil-type) HL-60 cells occlude the first constriction for only 4.3 msec before passaging through the subsequent constrictions with a median transit time of 3.3 msec. This method can provide insight into the viscoelastic nature of cells, and ultimately reveal the molecular origins of this behavior.

We demonstrate a microfluidics-based assay to measure the timescale for cells to transit through a sequence of micron-scale constrictions.

Here we detail the design, fabrication, and use of a microfluidic device to evaluate the deformability of a large number of individual cells in an ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

Electrotaxis Studies of Lung Cancer Cells using a Multichannel Dual-electric-field Microfluidic Chip

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of physiological dcEF in guiding cell movement during embryo development, cell differentiation, and wound healing has been demonstrated in many studies. By applying microfluidic chips to an electrotaxis assay, the investigation process is shortened and experimental errors are minimized. In recent years, microfluidic devices made of polymeric substances (e.g., polymethylmethacrylate, PMMA, or acrylic) or polydimethylsiloxane (PDMS) have been widely used in studying the responses of cells to electrical stimulation. However, unlike the numerous steps required to fabricate a PDMS device, the simple and rapid construction of the acrylic micro&#64258;uidic chip makes it suitable for both device prototyping and production. Yet none of the reported devices facilitate the efficient study of the simultaneous chemical and dcEF effects on cells. In this report, we describe our design and fabrication of an acrylic-based multichannel dual-electric-field (MDF) chip to investigate the concurrent effect of chemical and electrical stimulation on lung cancer cells. The MDF chip provides eight combinations of electrical/chemical stimulations in a single test. The chip not only greatly shortens the required experimental time but also increases accuracy in electrotaxis studies.

Many microfluidic devices have been developed for use in the study of electrotaxis. Yet, none of these chips allows the efficient study of the simultaneous chemical and electric-field (EF) effects on cells. We developed a polymethylmethacrylate-based device that offers better-controlled coexisting EF and chemical stimulation for use in electrotaxis research.

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Polydimethylsiloxane-polycarbonate Microfluidic Devices for Cell Migration Studies Under Perpendicular Chemical and Oxygen Gradients

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under combinations of chemical and oxygen gradients. Both chemical and oxygen gradients can greatly affect cell migration in vivo; however, due to technical limitations, very little research has been performed to investigate their effects in vitro. The device developed in this research takes advantage of a series of serpentine-shaped channels to generate the desired chemical gradients and exploits a spatially confined chemical reaction method for oxygen gradient generation. The directions of the chemical and oxygen gradients are perpendicular to each other to enable straightforward migration result interpretation. In order to efficiently generate the oxygen gradients with minimal chemical consumption, the embedded PC thin film is utilized as a gas diffusion barrier. The developed microfluidic device can be actuated by syringe pumps and placed into a conventional cell incubator during cell migration experiments to allow for setup simplification and optimized cell culture conditions. In cell experiments, we used the device to study migrations of adenocarcinomic human alveolar basal epithelial cells, A549, under combinations of chemokine (stromal cell-derived factor, SDF-1&#945;) and oxygen gradients. The experimental results show that the device can stably generate perpendicular chemokine and oxygen gradients and is compatible with cells. The migration study results indicate that oxygen gradients may play an essential role in guiding cell migration, and cellular behavior under combinations of gradients cannot be predicted from those under single gradients. The device provides a powerful and practical tool for researchers to study interactions between chemical and oxygen gradients in cell culture, which can promote better cell migration studies in more in vivo-like microenvironments.

The control of chemical and oxygen gradients is essential for cell cultures. This paper reports a polydimethylsiloxane-polycarbonate (PDMS-PC) microfluidic device capable of reliably generating combinations of chemical and oxygen gradients for cell migration studies, which can be practically utilized in biological labs without sophisticated instrumentation.

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under ...

A Simple and Scalable Fabrication Method for Organic Electronic Devices on Textiles

Today, wearable electronics devices combine a large variety of functional, stretchable, and flexible technologies. However, in many cases, these devices cannot be worn under everyday conditions. Therefore, textiles are commonly considered the best substrate to accommodate electronic devices in wearable use. In this paper, we describe how to selectively pattern organic electroactive materials on textiles from a solution in an easy and scalable manner. This versatile deposition technique enables the fabrication of wearable organic electronic devices on clothes.

In this paper, we present a protocol to selectively deposit organic materials on textiles, which allows for the direct integration of organic electronic devices with wearables. The fabricated devices can be fully integrated in textiles, respecting their mechanical appearance and enabling sensing capabilities.

Today, wearable electronics devices combine a large variety of functional, stretchable, and flexible technologies. However, in many cases, these devices ...

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...