A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Podsumowanie

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

Streszczenie

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

Wprowadzenie

Caenorhabditis elegans has proved to be a powerful model organism to study cell biology, aging, development biology, and neurobiology. Advantages such as its transparent body, short life cycle, easy maintenance, a defined number of cells, homology with several human genes, and well-studied genetics have led to C. elegans becoming a popular model both for fundamental biology discoveries and applied research^1,2. Understanding cell's biological and developmental processes from repeated long-term observation of individual animals can prove to be beneficial. Conventionally, C. elegans is anesthetized on agar pads and imaged under the microscope. Adverse effects of anesthetics on the health of animals limit the use of anesthetized animals for long-term and repeated intermittent imaging of the same animal^3,4. Recent advances in microfluidic technologies and their adaptation for anesthetic-free trapping of C. elegans with negligible health hazards enable high-resolution imaging of the same animal over a short and long period of time.

Microfluidic chips have been designed for C. elegans'⁵ high throughput screening^6,7,8, trapping and dispensing⁹, drug screening^10,11, neuron stimulation with high-resolution imaging¹², and high-resolution imaging of the animal^12,13,14. Ultra-thin microfluidic sheets for immobilization on slides have also been developed¹⁵. Long-term studies of C. elegans have been performed using low-resolution images of animals growing in liquid culture to observe growth, calcium dynamics, drug effects on their behavior^16,17,18,19, their longevity, and aging²⁰. Long-term studies using high-resolution microscopy have been carried out to assess synaptic development²¹, neuronal regeneration²², and mitochondrial addition²³. Long-term high-resolution imaging and tracing of cell fate and differentiation have been done in multichannel devices^24,25. Several cellular and sub-cellular events occur over the time scales of several hours and require trapping the same individual at different time points during their development to characterize all intermediate steps in the process to understand cellular dynamics in vivo. To image biological process such as organogenesis, neuronal development, and cell migration, the animal needs to be immobilized in the same orientation at multiple time points. We have previously published a protocol for high-resolution imaging of C. elegans for over 36 h to determine where mitochondria are added along the touch receptor neurons (TRNs)²³.

This paper provides a protocol for establishing a microfluidics-based methodology for repeated high-resolution imaging. This device, with a single flow channel, is best suited for repeated imaging of a single animal per device. To improve throughput and image many animals at once, multiple devices could be connected to the same pressure line but with separate three-way connectors controlling a single animal in each device. The design is useful for studies that demand high-resolution time-lapse images such as post-embryonic developmental processes, cell migration, organelle transport, gene expression studies, etc. The technology could be limiting for some applications such as lifespan and aging studies that require parallel growth and imaging of many late-stage animals. Polydimethylsiloxane (PDMS) elastomer was used for fabricating this device due to its biostability²⁶, biocompatibility^27,28, gas permeablility^29,30, and tunable elastic modulus³¹. This two-layer device allows the growth of animals with continuous food supply in a microfluidic channel and the trapping of individual C. elegans via PDMS membrane compression using nitrogen gas. This device is an extension of the previously published device with the advantage of growing and imaging the same animal in the microchannel under a continuous food supply³. The additional isolation membrane network and a 2 mm wide trapping membrane enable efficient immobilization of developing animals. The device has been used to observe neuronal development, vulval development, and dendritic arborization in sensory PVD neurons. The animals grow without adverse health effects in the device and can be repeatedly immobilized to facilitate imaging sub-cellular events in the same animal during its development.

The entire protocol is divided into five parts. Part 1 describes device fabrication for the growth and imaging chip. Part 2 describes how to set up a pressure system for the PDMS membrane deflection to immobilize and isolate individual C. elegans. Part 3 describes how to synchronize C. elegans on a nematode growth medium (NGM) plate for device imaging. Part 4 describes how to load a single animal in the device and grow the animal inside the microfluidic device for several days. Part 5 describes how to immobilize an individual animal at multiple time points, capture high-resolution images using different objectives, and analyze the images using Fiji.

Protokół

1. Fabrication of growth and imaging device

1. SU8 mold fabrication
   1. Design patterns 1 (flow layer) and 2 (control layer) using rectangular shapes in a word processing software (or a computer-aided design CAD software) and print the photomasks with the help of a laser plotter with a minimum feature size of 8 µm on polyester-based film (Figure 1).
   2. Cut silicon wafers in 2.5 cm × 2.5 cm pieces and clean them with 20% KOH for 1 min. Rinse the wafers in deionized (DI) water. Use one wafer each for the flow and the control layer.
      CAUTION: KOH is corrosive and should be handled with care.
   3. Dry the pieces with 14 psi compressed nitrogen gas followed by dehydration on a hot plate at 120 °C for 4 h. Before proceeding to the next step, cool the two pieces down to room temperature.
   4. Take one of the silicon pieces and put it on the chuck of a spin coater and turn on the vacuum to hold the wafer in place. On the silicon piece, put ~20 µL of hexamethyldisilane (HMDS) and coat it using the spin coater at 500 rotation per min (rpm) for 5 s followed by 3,000 rpm for 30 s.
      CAUTION: This step should be performed in yellow light. Do not use white light in the room.
   5. To get a uniform photoresist thickness of ~40 µm (specific to the flow layer; suitable for imaging early larval stage 1 to stage 3 (L1 - L3) animals), coat the silicon wafer with ~1.5 mL of negative photoresist-1 using a spin coater at 500 rpm for 5 s followed by 2,000 rpm for 30 s.
   6. Repeat steps 1.1.4 and 1.1.5 with the second wafer to obtain a uniform photoresist thickness of ~40 µm specific to the control layer.
   7. Alternatively, to increase the thickness of the flow layer to ~80 µm for older animals, coat silicon wafers with ~1.5 mL of negative photoresist-2 using the spin coater at 500 rpm for 5 s followed by 2,000 rpm for 30 s. This thickness is suitable for L3 stage to adult animals.
      CAUTION: Hold the silicon wafers by their sides to avoid any damage to the spin-coated layers. Keep white lights turned off during this step.
   8. Bake the photoresist-coated silicon pieces (for flow and control layers) on a hot plate at 65 °C for 1 min followed by 95 °C for 10 min. Cool the baked pieces to room temperature.
      NOTE: The baked silicon pieces can be stored for one day before proceeding to the next step. Store in the dark and do not expose to white light.
   9. Put soft-baked silicon pieces on the exposure stage of the UV illuminator with the photoresist-coated surface facing the UV lamp. Expose the two pieces separately to UV for 15 s, using a 200 W lamp, through a photomask with patterns 1 and 2 to get flow and control layers, respectively.
      CAUTION: Wear safety goggles and avoid direct exposure to UV light. Do not turn on white light in the room during this stage.
   10. Bake the two exposed silicon pieces with coated layer facing up, at 65 °C followed by 95 °C for 1 min and 10 min respectively. Cool the pieces to room temperature before proceeding to the next step.
   11. Develop the patterns by soaking the silicon pieces in the photoresist developer solution (1:3 dilution of the developer in isopropanol) for 20 min. Once the pattern is visible, rinse the pieces with pure iso-propyl alcohol (IPA) and gently blow dry using nitrogen gas (14 psi).
       CAUTION: Use a well-ventilated environment for this chemical treatment to avoid human exposure. Use white light only after features are developed and rinsed with IPA.
   12. Keep the silicon pieces in a desiccator with the coated surface facing up. Expose the pieces to silane vapors by pouring 50 µL of pure trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane on a small plastic cup or a glass slide. Place the cup/slide inside a desiccator and incubate for 2 h.
       CAUTION: Avoid direct exposure to silane vapor. Always use a sealed chamber for silane vapor treatment.
       NOTE: If necessary, the developed silicon pieces can be stored for 1-2 days before proceeding to the next step.
2. PDMS chip fabrication
   1. Make PDMS in a plastic cup by mixing the elastomer base with the curing agent in a 10:1 ratio. Mix the contents well by stirring constantly for 3 min. The mixing will create a lot of air bubbles in the PDMS mix.
   2. Degas the PDMS mix in a desiccator for 30 min to remove all air bubbles.
      CAUTION: Ensure air bubbles are removed from the PDMS mix before it is poured on the features since bubbles can cause defective and non-functional devices.
   3. Place the silicon wafers with the control layer (pattern 2) in a Petri dish. Gently pour a 5 mm thick PDMS mix layer on the silicon piece avoiding any bubble formation.
   4. Degas the PDMS mix in a desiccator to remove additional bubbles that are formed during PDMS pouring process.
   5. Place the silicon wafer with flow layer (pattern 1) on the spinner chuck applying 200-500 mTorr vacuum pressure to hold the wafer. Pour ~1 mL of PDMS on the silicon wafer and coat it using a spin coater at 500 rpm for 5 s followed by 1,000 rpm for 30 s to get an ~80 µm thick layer.
   6. Bake the two silicon wafers with the spin coated PDMS and poured PDMS layers at 50 °C in a hot air convection oven for 6 h. After baking, wait for the pieces to cool down at room temperature.
   7. Cut the 5 mm thick PDMS layer from the silicon piece around the control layer (pattern 2) using a sharp blade and peel it off from the silicon substrate.
   8. Punch two holes of ~1 mm diameter using a Harris puncher at the reservoir of the PDMS block to connect the immobilization channel and isolation channel inlets to the gas lines for PDMS membrane deflections.
   9. Place the silicon piece with the spin coated PDMS layer on the pattern 1 (flow layer), with the PDMS-coated surface facing up, on a plastic tray. Keep the punched PDMS block with pattern 2 (control layer) on the tray with molded side facing up.
   10. Keep the plastic tray inside a plasma cleaner and expose the two PDMS surfaces to 18 W air plasma for 2 min under low vacuum (200-600 mTorr). Apply vacuum until the chamber turns bright violet. Perform this step under low light to see the plasma color change.
   11. Take out the two plasma-treated blocks and gently bind the blocks by pressing the plasma-treated surfaces of patterns 1 and 2 together. Bake the bonded patterns at 50 °C for 2 h in a hot air convection oven.
   12. Take the bonded device out of the oven. Cut the bonded device out of silicon wafer with pattern 1 and pattern 2, and punch holes in the inlet and outlet reservoirs of the flow layer using the Harris puncher.
   13. Place the bonded PDMS block with the flow layer facing up on a plastic tray. Keep a clean cover glass (#1.5) on the same tray. Expose the blocks and the cover glass to 18 W air plasma for 2 min. Adjust vacuum pressure to see a violet chamber.
       NOTE: This step should be done in low light to see the plasma color change. To clean the cover glass, wash it with IPA and blow dry using nitrogen gas at 14 psi.
   14. Place the plasma exposed PDMS block on top of the cover glass and bake the bonded structure in an oven at 50 °C for 2 h. Store the device in a clean chamber for any future experiment.

2. PDMS membrane priming

1. Take the device and put it on a stereomicroscope and attach the tubings. Connect micro flex tube (inner diameter ~5 mm, outer diameter ~8 mm) to a compressed nitrogen gas line on one end. Connect a three-way connector on the other end. Tubes 1 and 2 of the three-way connectors will be connected to the trap and isolating membranes, respectively.
2. Connect two micro flex tubes (inner diameter ~1.6 mm, outer diameter ~5 mm) to the two outlet ports of the three-way stopcock. Connect the other end of the two tubes to an 8 mm long 18 G needle.
3. Fill the flow layer with M9 buffer using a micropipette through the inlet port. Fill both tubes with DI water through the end connected to the needle. Insert the two needles into the punched holes, connecting the isolating and trapping membrane, respectively.
4. Open the nitrogen gas regulator at 14 psi and turn the three-way valve from tube 1 to push the water into the device through the microfluidic channels in the control layers namely, the trap and isolation membranes.
5. Wait until water fills the channel without the presence of any air droplet in both channels. Once the channels are completely filled with water, the channels are considered to be primed.
6. Release the pressure using the three-way stopcock once the channels are filled with water and primed. Priming can lead to bubbles in the flow layer, remove the bubbles by flowing additional media through the flow channel.

3. C. elegans maintenance and synchronization

NOTE: C. elegans strains: The study used following transgenes PS3239 (dpy-20(e1282) syIs49 IV [MH86p(dpy-20(+) + pJB100(ZMP-1::GFP)]) for vulval development³², jsIs609 (mec7p::MLS(mitochondrial matrix localization signal)::GFP)³³ for touch receptor neuron (TRN) development and mitochondria transport imaging, and wdIs51(F49H12.4::GFP + unc-119(+)) to track PVD development³⁴. Standard C. elegans culture and maintenance protocol was followed³⁵.

1. Grow C. elegans on the nematode growth medium (NGM) Petri plates with E. coli OP50 as the food source at 22 °C. Maintain the C. elegans strains by repeatedly transferring a few hermaphrodites or chunking a small amount of agar with a few animals to a new NGM plate with an OP50 lawn.
2. After 3-5 days, check the NGM plate for animal growth and C. elegans eggs. Collect and transfer approximately 30 eggs from a plate to a fresh NGM plate with OP50 lawn.
3. To synchronize animals for imaging, transfer all unhatched eggs from the plate every 2 h to a fresh plate and maintain them at 22 °C. Approximately 15-20 eggs will hatch on each plate.
4. Pick the animals between 14-16 h and 28-30 h after hatching for larval 2 (L2) and larval 3 (L3) stages, respectively. Transfer the animals to the microfluidic device for imaging. Add food supply as described in the next section to maintain the animal inside the device for long-term growth and imaging experiments.

4. C. elegans growth inside the growth and imaging microfluidic device

1. Mount a growth and imaging microfluidic device on an inverted microscope and view pattern 2, after connecting the isolation membranes and immobilization membrane, at low magnifications (4x or 10x). Ensure that the channels are filled with clean distilled water.
2. Prepare a fresh 1 L solution of S medium using 10 mL of 1 M potassium citrate pH 6.0, 10 mL of trace metals solution, 3 mL of 1 M CaCl₂, 3 mL of 1 M MgSO₄. Prepare the solution under sterile conditions. Do not autoclave the S medium.
3. Fill the flow channel with growth media (S medium) 10 min before the experiment. Avoid any air bubbles in the flow channel. Flow additional medium if needed to remove air bubbles.
4. Pick a single animal from the required developmental stage from an NGM plate in 10 µL of S medium using a micropipette and push the animal into the flow channel through the inlet hole.
5. Monitor the animal position in the flow channel using a low magnification objective. Flow additional medium through the inlet or outlet to push the animal and position it within the flow channel restricted between the two isolation membranes.
6. Open the three-way stopcock to apply 14 psi pressure in the isolation channels and push the membranes down into the flow channel.
   NOTE: The membrane partially seals the flow channel and restricts animal movement to the region between the two isolation membranes.
7. Use a single colony of E. coli OP50 from a streaked plate to inoculate 250 mL of L Broth (2.5 g bacto-tryptone, 1.25 g bacto-yeast, 1.25 g NaCl in H₂O). Grow the inoculated culture overnight at 37 °C.
8. Aliquot 500 µL of OP50 culture into 1.5 mL sterile centrifuge tubes and store the stock and the aliquot for 2 weeks at 4 °C.
9. Pellet down OP50 culture by centrifugation at 1.3 x g for 5 min. Dissolve the pellet with 1 mL of fresh S medium (0.5x dilution) and store it at room temperature for 3-4 days. Use this diluted OP50 to feed C. elegans inside microfluidic devices.
10. Leave a drop of S medium on top of the inlet and outlet reservoirs to reduce evaporation of the S medium in the flow channel.
11. Take diluted OP50 solution in a 10 µL micropipette. Remove the micro tip filled with the OP50 solution from the pipette and press-fit the tip into the inlet reservoir. Then place another micropipette tip with 10 µL of food solution and insert it into the outlet reservoir.
12. Seal the tip head applying pressure using a finger to ensure continuity of food solutions without any air gap. Change the micropipette tip with OP50 solution every day that is not older than 3-4 days.
13. Fill an additional 20-30 µL of food solution in both the tips. Add or remove food solution to and from the micropipette tip to adjust the gradient to push the animal in the flow channel and to adjust the position of the animal under the trapping membrane for imaging.
14. Monitor the flow of bacteria to ensure it is continuous through the partially closed isolation membranes in the flow channel using bright-field images.
    ​NOTE: In absence of bacteria flow, animals might eat the available bacteria in the flow channel between the two isolation membranes. In case of no bacteria or no flow present in the channel, replace the two micropipette tips at the inlet and outlet with new pipette tips filled up with freshly prepared food supplies.

5. C. elegans immobilization and imaging

1. C. elegans immobilization under the trapping membrane
   1. Locate a single animal in the growth channel at low magnifications. Adjust the food solution heights in the two micropipette tips connected to the inlet and outlet reservoirs. Push the animal in the required direction using the hydrostatic pressure differences in the flow channel.
   2. Position the animal at the center of the trapping membrane and monitor its swimming behavior using a low magnification objective (4x).
   3. Turn the three-way stopcock to increase the pressure in the trap channel slowly. Immobilize the animal under the trap membrane in a straight posture along the growth channel boundary wall.
      CAUTION: Avoid trapping the animal across the flow channel in a Z or U bend position. This causes the animal body to be squeezed with greater pressure and causes permanent damage to its health.
2. C. elegans imaging and release from the membrane
   1. Carry and load the device on a microscope stage. Set up an inverted microscope at the desired imaging settings with all the necessary optical components (objective, light source, fluorescence filters, and a detector) for high-resolution bright field, differential imaging contrast (DIC), or fluorescence imaging (Supplementary Figure 1).
   2. Acquire a single or several time-lapse fluorescence images to capture cellular and sub-cellular events.
   3. Acquire fluorescence images of C. elegans neurons as a function of their development using a 60x, 1.4 numerical aperture (NA) objective, a 488 nm wavelength laser with 7%-10% laser power, a CCD camera, on a spinning disc microscope. Perform time-lapse imaging using the software provided with the microscope and acquire images at a speed of 4 frames per second (Supplementary Figure 1).
   4. After the acquisition of images, release the trapping pressure and monitor the locomotion of the animal at low magnifications - 4x and 10x. Keep the animal restricted within the region defined by isolating membranes (kept under 14 psi all through the experiment).
   5. Adjust the volume of food solution in the two micropipette tips to continue a slow gravity-driven food flow in the growth channel. This food flow is obtained by adjusting the levels of the food solutions in micropipettes at the inlet and the outlet (typically 1-5 mm height difference).
   6. Visualize the flow under a bright-field from the flow pattern of the bacteria in the growth channel.
      NOTE: In the absence of flow, the animal eats the available OP50 bacteria, the channel appears clear, and the animal starves over a few hours. A starved animal typically develops high autofluorescence, easily observable when acquiring time-lapse fluorescence images. Such events have detrimental effects on animal physiology and were avoided in the experiments.
   7. Repeat steps 5.2.1-5.2.3 after a predetermined time interval to acquire fluorescence/ DIC/bright-field images of the same individual at multiple time points.
   8. After the images are acquired at multiple desired time points from the same individual animal, release the isolation and trapping pressure. Flush the channel with M9 buffer and push the animal through the inlet reservoir. Recover the animal from the reservoir and place the animal on a fresh NGM plate for further health monitoring.
   9. Flush the channel with M9 buffer a few times to remove bacteria. Rinse the flow channel with 70% ethyl alcohol (diluted in distilled water). Dry the channels by pushing air using a syringe. Store the device in a dry dust-free place for repeated use in the future.
3. Image analysis and statistics
   1. Use FIJI ImageJ software for tiff image analysis. Open tiff images of the PVD time-lapse images from the wdIs51 animals in Fiji ImageJ. Extract the best planes from the stack of z-planes showing the primary processes by selecting Image tab > Stacks > Z project.
   2. Screen the entire series of images and find specific image frames that successfully cover the entire neuronal processes using overlapping sections of the animal image frames. Select Plugin > Analyze > Cell Counter from the FIJI toolbar. This will open a window for numbering different branches and keeping a count of each selected neuron.
   3. Select Initialize > Counters > Type1, then mark the secondary branches. Then select Type 2 and mark Quaternary, click Results window showing counts of all branches (Supplementary Figure 2). This method will display the branches that are already counted.
   4. Open the next overlapping image and count the remaining branches. This process will prevent double counting and ensure every process is counted. Identify and count the total number of primary branches present in the early larval stages of C. elegans. Perform similar analysis for the images for secondary and quaternary processes in older animals.
      NOTE: Adults show many processes that innervate in body wall muscles and can be difficult to count. Ensure that every process is counted only once.
   5. Calculate the distance between the PVD cell bodies in the wdIs51 animals by drawing a segmented line from the PVD cell body (CB) to PVC CB. For larval 4 (L4) stage animals, the cell bodies are far apart and are not in the same frame when imaged with a 60x objective.
   6. Select overlapping images from stack to cover the entire animal length from head-to-tail using Image tab > Stacks > Z project from ImageJ. Draw segmented lines along the neuronal process between the PVD and PVC CBs.
   7. Measure the lengths for all the segmented lines using ImageJ > Analyze > Measure. Add the lengths for all the segments to calculate the total distance between PVD and PVC CBs for each time point.
   8. For neuron length of TRNs in jsIs609 animals, load the images in Fiji. Draw a segmented line along the posterior lateral microtubules (PLMs; visible with soluble GFP) from the cell body to the centroid of the first mitochondrion. Calculate the length of the line for the first distance value.
   9. Draw a segmented line from the center of the first to the second mitochondrion. Repeat this process for each pair of mitochondria till the end of neuronal process length. Add all the lengths to calculate the total neuronal process length.
   10. For the cell lineage analysis, load all the vulval images in Fiji and extract the best focus image of the cells from the time-lapse images of multiple z-planes.
   11. Represent the data as mean ± standard error of the mean (SEM). Calculate statistical significance using one-way ANOVA for more than two samples or a two-sample t-test for a pair of samples. Denote significance by p-value < 0.05 (*), p-value < 0.005 (**), and p-value > 0.05 (ns, not significant).

Wyniki

Device characterization: The growth and imaging device consists of two PDMS layers bonded together (Figure 1) using irreversible plasma bonding. The flow layer (pattern 1) which is 10 mm in length and 40 µm or 80 µm in height allows us to grow the animal in liquid culture (Figure 1A). The trapping layer (pattern 2) has a 2 mm wide membrane (Figure 1B) for immobilizing the animal for high-resolution imaging...

Dyskusje

In this paper, a protocol for fabrication and use of a simple microfluidic device for growing C. elegans with constant food supply and high-resolution imaging of a single animal during its development has been described. This fabrication process is simple and can be done in a non-sterile environment. A dust-free environment is critical during fabrication steps. The presence of dust particles would lead to improper contact between the two bonding surfaces, resulting in poor bonding and leakage of the device durin...

Materiały

Name	Company	Catalog Number	Comments
18 G needles	Sigma-Aldrich, Bangalore, India	Gauge 18
3-way stopcock	Cole-Parmer	WW-30600-02	Masterflex fitting with luer lock
CCD camera	Andor Technology	EMCCD C9100-13no
Circuit board film	Fine Line Imaging, Colorado, USA		The designs are printed with 65,024 dots per inch (DPI)
Convection Oven	Meta-Lab Scientific Industries, India	MSI-5
Coverslips	Blue stat microscopic cover glass		22mm x 10Gms
Ethanol	Hi media
Harris uni-core puncher 1mm	Qiagen	Z708801
Hexamethyldisilazane	Sigma-Aldrich, Bangalore, India	440191
Hot plate	IKA	RCT B S 22
Isopropanol	Fisher Scientific	26895
KOH	Fisher Scientific
Laser Scanning Microscope	ZEISS	LSM 5 LIVE
Micropipette tips	Tarsons		0.5-10 µL micropipette tips are used for food supply
Negative Photoresist-1	Microchem	SU8-2025	http://www.microchem.com/Prod-SU82000.htm
Negative Photoresist-2	Microchem	SU8-2050	http://www.microchem.com/Prod-SU82000.htm
Nitrogen gas	Local Supplier	Commercial nitrogen gas	Cylinder volume of 7 cubic meter
PDMS (Curing solution)	Dow Corning Corporation, MI, USA	Sylgard curing solution	curing agent
Petri plates	Praveen Scientific Corporation
Plasma cleaner	Harrick Plasma, NY, USA	PDC-32G
Razor and blades	Lister surgical Blade
Silicon Elastomer (Base)	Dow Corning Corporation, MI, USA	Sylgard 184 base	elastomer base
Silicon tubes	Fisher Scientific		Plastic tubes with the inner diameter 1.59 mm and the outer diameter 3.18 mm
Silicon wafer	University Wafer, MA, USA	[100] orientation, 4-inch diameter	Small pieces (2 mm × 2 mm) were cut from 100 mm diameter wafer
Spin Coater	SPS-Europe B.V., The Netherlands	SPIN 150
Spinning Disk microscope	Perkin Elmer ultra-view VOX system	CSU-X1-A3 N	The system was equipped with four (405/488/561/640 nm) lasers and controlled with the Volocity software package.
SU8 developer	Microchem, MA, USA	SU8 Developer
Trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane	Sigma-Aldrich, Bangalore, India	448931	Trichloro (1H, 1H, 2H, 2H-perfluorooctyl) silane vapor is toxic
UV lamp	Oriel Instruments, Bangalore, India		200 Watt and collimated UV light source
Volocity software	Perkin-Elmer		Image analysis