AsianScientist (Jun. 27, 2016) – So, it’s a Friday night and you’ve been hard at work at the bench all week. You’re feeling exhausted and absolutely frustrated. Some experiments have left you scratching your head and wondering if you’re really cut out for science.
Cells getting lysed when you rinsed them with phosphate buffered saline? Random bands appearing on your DNA agarose gel? Huge error bars on your qPCR run? What are the odds of everything going wrong at once? Well, it sounds like you need to break something—I mean, take a break or something.
And while you sip on a cup of hot coffee or down a pint of cold beer, read these ten hacks that might make your lab experience a little more pleasant.
- Treat unlabeled reagents like toxic waste
Perhaps your colleague told you that the unlabeled tube in the biosafety cabinet contains phosphate buffered saline. The liquid inside the tube is clear and odorless as it should be. Discard it anyway. Sometimes, the most innocuous items left lying around the lab can ruin an entire week of work and planning.
If you didn’t leave it there yourself and it doesn’t have a label, treat it like toxic waste and throw it out.
- Spike your PCR
If your PCR is giving you unexpected outcomes when you eventually run the DNA agarose gel, it could be secondary structure that’s interfering with primer annealing and/or amplification.
DMSO and betaine monohydrate are just some of the additives that have been known to help with PCR problems and they work especially well to untangle issues with GC-rich templates. Like all optimization steps, you’ll need to do some reading and reagent titrations to find the best conditions for your PCR run.
- Enhance SYBR security
When making up your qPCR master mix, instead of combining water, primers and SYBR green altogether, mix water with your template cDNA in one tube and SYBR green with primers in another so you now have two ‘half-master mixes.’
For each 20µL reaction, make up the half-master mixes as follows: 2µL of cDNA with 7.2µL of water in tube A, 0.8µL of 10µM primers with 10µL of SYBR green in tube B.
This way, you won’t have to pipette small volumes of less than two microliters into each well which drastically reduces pipetting error, thereby shrinking your qPCR triplicate error bars significantly.
- Tame the wild wild West
We all have encountered Western blots that look like abstract art. Sometimes, the bands come out clearly but they’re slanted, ‘frowning’ or ‘smiling’ as if they’re privy to some jest at your expense. This usually happens when the salt concentration in your sample buffer is too high, or you’re loading too much protein into the wells.
If a noisy background is your bugbear, try more stringent blocking conditions such as a higher percentage of BSA or goat serum. Extending the number of blocking hours also helps, and if you decide to do it overnight, keep the membrane and blocking solution on a shaker at 4 °C. Adding 0.1 % Tween-20 to your washing solutions after primary and/or secondary antibody incubation also makes for a ‘cleaner’ blot.
- Get your fix(ative) fast
If you’ve ever had to make up four percent paraformaldehyde in phosphate buffered saline, you’ll know that the white fixative powder takes forever to dissolve even if you’re using saline heated to 60 °C.
Here’s a tip: add a few drops of 1 N sodium hydroxide to the beaker and watch the powder transit seamlessly into solution within five minutes. You’ll have to adjust the pH back to 7.4 after that with hydrogen chloride, but that can be done really quickly.
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