Step aside DJs – biochemists are the real master mixers! And the master mix isn’t the last of our tricks for similar-but-different-but-lots of samples! Often the lab you have to set up a bunch of reactions that are almost the same – they just differ in 1 or 2 things – like you want to radiolabel different RNAs and all the things you need for the labeling (water, buffer, kinase, ATP) are the same, but the RNAs are different. Since all the reactions are the same except for that thing I can prepare a “master mix” of all the “same stuff” – so that I only have to add 1 thing per different thing instead of 4
It’s kinda like if a company has a form they need all their employees to sign. If the company had to type up the form individually for each person, that would take a really long time – and each time introduces the chance for the company to make a typo – and it can be pretty easy to get exhausted and or boredly mind-wandery, etc. after typing the same thing out over and over. The lab version of this is what I call “pipetting apathy”
So, the company instead makes 1 form and just leaves room for the employee to sign. If there are a couple of things that can vary (e.g. maybe they need to print their name and sign, and/or write down the date) they can leave space for those too.
Then they only have to type it up once, can distribute this single document to all their employees, and don’t have to worry that they accidentally gave 1 employee a form where they left out the “not” before “liable” – getting their employee to sign that the company *is* liable for damages…
The company doesn’t just print out the number of copies they need – they print extra so that if someone messes up their form (pipets the wrong thing or wrong volume, etc.) or loses the form (drops a tube or something) they don’t have to type up a whole new form.
When it comes to pipetting the situation’s a bit more complicated too because instead of having defined sheets, everything is mixed together, so it’s more like ladling punch. Each time you ladle, a bit gets stuck on the ladle, some may spill, some evaporates, etc. so you want to make a bit more than you need (e.g. if you have 20 guests & you want each guest to get exactly 1 cup of punch, you don’t want to make exactly 20 cups-worth of punch or the last person will get jipped.
So when making master mixes, you always want to prepare for more reactions than you actually need – because every time you pipet, some gets stuck to the pipet tip, some can evaporate, etc. & you want to make sure you have enough. If all the stuff’s cheap, it’s good to make enough for a little more than 1 extra in case you mess up on 1 reaction you have enough to redo it. But when the reagents are expensive and/or radioactive, I just make enough extra to account for some minor losses.
If it’s something you do a lot and/or there are lots of components with decimals and stuff, you can even make a spreadsheet where you have a “per reaction” column and then a “total column” with the amounts you get if you multiply it by a “conversion factor” you can change (if you’re using excel, use F4 after you click on the well you want it to multiply by (the conversion factor so if you try to copy the formula it still knows where to look).
Another think to watch out for – remember that the total reaction volume is the master mix PLUS the unique thing. So for a 50uL reaction volume, remember that you don’t add 50uL of it to your thing.
So, when I was radiolabeling RNA the other day (more here: http://bit.ly/2lb0O9U I I premixed the buffer, water, hot ATP, & PNK – and added 48ul of that directly to 2uL of RNA to get to the final reaction volume of 50uL. I do this type of “master mix” thing a lot. It’s a big thumb-saver & tip-saver and it helps ensure that all the tubes are getting the same amount of everything and you don’t accidentally skip one of the components for one tube, etc.
A couple of other huge time-savers and consistency-enforcers which you can use with master mixes or with single things are the multichannel pipette – allows you to pipet the same amount of liquid into multiple tubes AT ONCE. And the repeater pipet – allows you to pipet the same amount of liquid multiple times, into whatever you want but NOT all at once.
With the multichannel you suck up (aspirate) once and dispense once – it’s just like as if you had a bunch of normal pipets taped together with 1 plunger. You suck up the amount you dispense and you dispense all at once. Like you go to the bank and get 8 $20 bills (or 12 if you have a 12-channel) and you call over your 8 friends and give them each a $20 at the same time.
I keep a PCR strip tube with SDS-PAGE sample buffer & 1 with urea-PAGE sample buffer so I can prep gel electrophoresis samples in PCR strip tubes when I have a lot.
With a repeater pipet you suck up once, but you suck up more than you plan to dispense each time and then each time you press down the plunger it releases a defined amount – for example, if you use a 10mL repeater pipet tip you can dispense 10 x 1mL portions, 20 x 0.5mL portions, etc. It’s more like you get a bunch of $20 bills from the bank and you can give them to whoever you want whenever you want but you can’t “break change” and give someone a $5 and you can only give them out 1 at a time.
It’s good for making aliquots of things that you want to freeze in “single-use” sizes to avoid unnecessary freeze-thaws (like competent cells, enzymes, etc.)
If you do want smaller volumes, there are different tip sizes for the repeater pipet and different whole pipets for the multichannels (and different tips to match). The repeater pipet & the multichannel have different strengths and weaknesses. A couple of weaknesses of the multichannel…
The tips are at fixed distances – they’re spaced so that they align with the well/tube spacing of blocks & PCR tube strips, but sometimes you want to pipet into things that aren’t spaced just like that. Sometimes you can rearrange the tip rack to compensate (e.g. I remove every other row and use a 12-channel to pipet the 6-across way into the slots of this slot blot (a device I use for binding experiments).
And since the pipet tips each draw up from those fixed locations, you need the stuff you want them to suck to be spaced out (or spread out) too. You can either have the liquid in something like a tube strip or a deep-well block. When you’re using one of these you need to make more than you need because each well needs a little extra, etc.Or you can use a reagent reservoir which is like a little V-shaped basin (the V helps the liquid collect at the bottom of the V).
With the repeater pipet, a couple limitations are – although you only have to suck once, you have to dispense multiple times – you’re restricted to set volume amounts (also – on a practical note – the first lever press gets you to the start point (e.g. don’t trust the first dispensing -it’s just letting out the excess so you’re ready to go, so eject it into the tube you’re drawing from not one of the places you want to put it.)
Our repeater pipet lost its battery backing – so if you’ve seen it let me know! And speaking of seeing things,,, When you’re working in deep-well blocks it can be really hard to tell the wells apart and really easy to forget which well you just took things from or put things into. Color-coding to the rescue!
I use different colored sharpies to color code the block and make different rows & columns stand out. And if I have to pipet from one block into another, I color code them in the same scheme so I can say add blue to blue.
And speaking of coming to the rescue – time-savers save my brain cells because I have a bad habit of holding my breath when I concentrate & when I pipet things (I think my brain is trying to sabotage my experiment because that oxygen’s pretty important – and so’s sleep so sorry for poor formatting, etc. – I spent my day using the multichannel pipet a lot!)