This information is probably obvious to most people reading this site, however I felt it was important to include for those new to bench work-- particularly undergrads. I've also put this category unalphabetically at the top, so my undergrads read it first.

• Read the Material Safety Data Sheet (MSDS) for all chemicals/materials you use! The MSDS should come with the chemical, or be available on the manufacturer's website. The MSDS tells you great stuff such as, "don't inhale this" or "this is harmless." See below for a good example.
• METHANOL is nasty stuff. But because it sounds like ethanol (which is alcohol, which people drink) and often sits quietly near Ethanol, Acetone, and IPA, it seems that it is treated as harmless. Quoth Wikipedia, "Methanol is extremely toxic. If ingested, as little as 10ml can cause permanent blindness and as little as 60ml can result in death."

• [New] If you're trying the replicate the glass capillary-based hydrodynamic focusing devices made by the Weitz Lab, I've found that WPI 1.0/0.58mm OD/ID (#1B100) fit perfectly inside Vitrocell 1mm ID square capillaries (#8100-100).
• You can take advantage of the flexibility of PDMS (see Methods).

• Get ImageJ.
• PDMS loves to suck up molecules. If you use a low molecular weight fluorescent molecule, such as Rhodamine or FITC, it'll absorb into your channel walls extremely quickly and leave obnoxious background fluorescence. Use a higher molecular weight fluorescent molecule such as FITC-Dextran that doesn't absorb as easily.
• You can also passivate your channels with BSA (see Methods), to inhibit absorption by PDMS. However, I've found that weird crystals can form that clog <25micron wide channels.
• If you're seeing too much background light or fluorescence from tubing, stick a piece of electrical tape on the top of your PDMS device. It helps a lot.
• If you're doing any sort of fluorescent quantitative analysis (such as measuring a gradient), you need to perform a flat-field correction to make sure you're not simply measuring differences in excitation. Also, align the arc-lamp in your microscope system if you can. I usually use a drop of fluorescent solution on a slide, and under a cover glass for my flat field correction.
• If you're measuring fluorescence over time, make sure you correct for photo bleaching.
• MicroManager is an open source microscope control/automation package.
• Your eye can't see (or likely your monitor display) the entire range of intensity values your CCD can capture. To get the full range of data from your fluorescent images, adjust your exposure time such that the graph of intensity levels is as broad as possible without saturation. Sisi Chen adds, "if you want to take multiple images over time, you also must take care that your fluorescent exposure is not TOO high. Otherwise, the cells might experience UV-induced cytotoxicity over the course of your experiment."

• Food coloring is great for filling your devices, and taking pictures. However, it will dry up over time.
• Food coloring is filled with microscale fibers and stuff, and can easily clog devices. You can filter the junk out, but it turns the food coloring a dark brown color.
• You won't be able to see the colors in food coloring at very high magnifications (>20X or so). Use a fluorescent dye.

• If you need information on polymers (e.g. their solvents, non-solvents), check out the Polymer Handbook.
• [New] If you need information on chemicals, check out the CRC Handbook of Chemistry and Physics.
• A very extensive resource for cell and tissue engineering is the Khademhosseini Lab protocols. I highly recommend skimming them.

• Keep up on the latest PubMed results: PubCrawler.
• If you have a Mac, get Papers.
• Also awesome, Zotero.
• Whitesides' Group: Writing a Paper
• [New] Weitz Lab Guide to Writing a Good Paper

• I leave about 2mm around a hole I plan to punch.
• If you are having trouble getting AutoCAD to region some of your polylines, save your file as a DXF, import it into LinkCAD (for conversion from DXF->DXF), "Repair" all broken polylines, and open your repaired file in AutoCAD.
• Cheap masks: Fineline Imaging, Photoplot Store
• [New] Mask Cleaning: An effective way to clean your mask is to douse it in Acetone, let that puddle of acetone sit for a few seconds, wipe it up with a kimwipe in both directions. Follow up in the same way with IPA, and then DI water. I used to try to clean my masks by spraying them with acetone over a waste bottle, but letting the acetone sit and wiping it off is much more effective.

• Degassing: If you need to degas rapidly, you can put your gassy PDMS mixture in a centrifuge tube, and centrifuge for a minute at a high speed (7g). Voila, bubbles gone. Don't spin too long, you don't want to separate the curing agent from the base. Also, bear in mind, centrifuge tubes are much more expensive than our cheap plastic cups.
• [New] Degassing: If your PDMS is mostly degassed, but you need to get rid of the last air bubbles quickly, slighty blow on your PDMS, or blow it lightly with N2.
• Cleaning: The best way to clean PDMS before a strong plasma bond is with Scotch Tape. Just apply the tape and pull it off. Scotch tape. Not lab tape, not duct tape, not electrical tape, not IPA, not Acetone. Scotch tape. The plasma clean will do the rest
• Cleaning: If you need to do a more thorough clean, rinse with Acetone, then IPA, then DI water. Rinse, don't let the PDMS soak. PDMS will swell in those solvents.
• Bonding: I seem to get much better plasma when the slits in the Plasmod glass chamber are facing downward (I have in no way tested that systematically). Tanner Nevill makes an excellent counter-point: If something spills in the Plasmod, and the slits are faced downward, things could get sucked into the vacuum.
• Bonding: I plasma my PDMS and slide for 1 minute (I get asked that a lot).
• Bonding: After plasma bonding, let your device sit in the oven (~60C) for at least 20 minutes. It significantly strengthens the bond, particularly for PDMS-PDMS bonding. Higher temperatures and hot plates work too.
• Bonding: Bonding without plasma, only HCl (see Methods). I haven't actually tried it.
• Curing: PDMS does not generally cure on top of glue and leaves a nasty sticky film. It does cure on top of Elmer's glue and Hot glue, but does not on top of Loctite Super Glue, Krazy Glue, Norland 68 Optical Adhesive, Hardman General Purpose Epoxy.
• Curing: PDMS will cure on top of Scotch tape and electrical tape.
• Curing: Rapid and clean curing. I actually put the wafer on top of a 4" diameter acrylic circle.
• Curing: If for some reason your PDMS leaves uncured goo on your wafer, just try pouring another layer of PDMS+curing agent on your wafer. Usually that will cure the goo.
• Curing: Do not try going lower than 3:1 Base:Curing agent. There's too much curing agent, and it doesn't solidify.
• You cannot leach Polystyrene beads out of cured PDMS with Toluene. Trust me.
• If you're so inclined to get beads in your PDMS, you'll encounter problems mixing the bead solution and PDMS (the bead solution just breaks up into smaller droplets). I managed by: pouring PDMS base into a dish, pour your bead solution on top of the base, leave the dish in the oven until all the solution from your beads evaporate, then add your PDMS curing agent and mix vigorously.
• Punching: Good tools for punching holes in PDMS. We use .042x.031x1.5 304 SS TiN Coated Round Punches (CR0420315N19R4).
• Punching: You can also punch holes with grinded down/flattened syringe tips.
• [New] High Aspect Ratios: If you're having trouble molding high aspect ratio (i.e. tall and thin) structures, PDMS might not be your best material because these structures can collapse. Check out these papers and the references therein.

• Pumping with filters on your syringes will cause very inconsistent flow rates. I do not recommend it.
• The easiest way to get bubbles air out of a complex device (a device that traps air and bubbles) is to fill the device with DI water, plug all the inlets and outlets except for one, and flow (gently apply pressure to a syringe) into that inlet until all the gas diffuses out through the PDMS.
• Gravity driven pumping is much, much more steady than using syringe pumps. You have to make sure to get bubbles completely out of all your tubing and device first, otherwise they will completely stop all flow.
• Over long times, and at low flow rates, devices in the incubator can easily have the solution in them evaporate, killing your cells. Put your device slide in a petri dish, leave a little water in the dish (don't drown your device), and cover it.
• Cells will quickly settle at the bottom of a syringe. Avoid that during device loading.
• Inline bubble trap and avoiding bubble injection.
• Diameters of various syringes

• If you want to use the double gap in the Malvern Bohlin Gemini rheometer, you have to zero it using the cone-and-bob attachment with the collar on. If you forget the collar and try to zero your gap, the rheometer will get stuck in the lowered position. Turn off the machine, and locate the nut on the top center of the machine. You can turn that to manually crank up the rheometer and try again.

• [New] If you need one wafer, consider just ordering it from the Stanford Microfluidics Foundry to save time and money.
• SU-8 adheres extremely well to SU-8. If you have structures that are peeling off your wafer, try first curing a thin base layer of SU-8 on your wafer and patterning your features on top of that.
• [New] For SU-8 adherence, I've also had great success by dehydrating my wafers (120C, 5 min) and then spinning on a layer of OmniCoat.
• A 20 minute or so final bake of your wafer at 150C will significantly help the adhesion of your SU-8 features to your wafer.
• SU-8 2035 has a much lower viscosity and is much easier to handle than SU-8 2050, but can still make 50 micron high structures.
• Getting closer to 90degree side walls (see Methods).
• SU-8 does not reflow (I don't think any negative photoresist will).
• [New] If you want to increase the lifetime of your mold, a common technique is to make your final structure in SU8, mold it in PDMS and use that PDMS as your regular use mold. These papers will clarify that.

• You can use the laser to cut through a thin layer of PDMS, it just leaves ugly, burnt sidewalls.
• If the laser isn't cutting all the way through your acrylic, make sure the lens size is set to the appropriate setting in the Options (2.0" for us). Also, clean all three lenses (one on the left, two on the right) with lens paper and cleaner.
• If you want to cut tape or some thin film on top of a glass slide, and not damage the slide, use these settings: Red lines, Paper setting, -20% power, thickness 0.001".
• If you're having trouble getting the laser to cut all the way through your acrylic, restart the computer. It will recalibrate the laser.
• The spot size of the laser is on the order of 500microns.
• Trying to etch into (not cut through) acrylic in order to make channels doesn't work too well, because the laser pulses. You end up with a very bumpy and crappy etch.
• You can glue pieces of acrylic together with SuperGlue. However, the glue won't dry easily. Breathe on it to speed the drying. I'm not kidding.

• Advice for graduate and undergraduate students in the sciences