My thumbs got weary just seeing the experiments I planned out for today – lots of serial dilution-ing – thank goodness for master mixes and multichannel pipets! Oh – and my eyes would like me to please thank sharpies, cluster tubes, and color coding! Today – time- and thumb-saving tips from the bumbling biochemist! (especially for when you have serial dilutions and/or a lot of reactions to do that are mostly the same)
Let’s start with the Master Mix. In my first lab rotation (where you try out different labs before choosing) the postdoc I was working with used this term and I’d never heard it but then I learned that it’s actually really common. So what is it?
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, you can prepare a “master mix” of all the “same stuff” – so you 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.
For example, when I was radiolabeling RNA the other day (more here: http://bit.ly/radiolabelings) 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 and use the multichannel to add the buffer.
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).
True story: I was terrified of using the repeater pipet for several months and instead stuck to just pipetting over and over. This was because the first time I used it it was for pipetting tiny volumes into crystallography wells and sometimes, since the volumes were so low, I couldn’t tell whether anything had come out – and sometimes I accidentally double-pressed it. Thankfully, it is a lot easier and more comfortable with target volumes!
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). Keep an empty tip box to put the ones you remove in so you can still use them!
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). Even with the basin, you still want to make extra because the liquid will get spread out even in the area in-between where the tips will try to suck from. So you need to make sure there’s enough to suck up at each site the tips try to suck from.
Another word of warning with the multichannel – make sure that all the tips are stuck on evenly – sometimes, depending on how you apply pressure when loading, some of them will be a little looser and take up different amounts. Each time you suck up, look at the levels of liquid drawn up and make sure that they’re even in all of the tips before you complete the transfer.
And, final word of advice for the multichannel: if you don’t need all of the 12 or 6 or whatever tips, center the tips you’re using in the center of the pipet (so, for example, you want to do
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We also have a couple of super-multichannel pipettes. A couple are “Liquidators” which allow you to transfer 96 wells at once – major thumbsaver for preparing crystal screens! We have a manual one and – one of my favorite things in the lab – an automated one!! (I use this one to help me with adding buffer to all my wells before doing serial dilutions, which I will tell you about later). We also have a robot called a Mosquito which dispenses teeny-tiny amounts of liquid into crystallography trays. https://www.instagram.com/p/BqaoNJchr-O/
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.)
One of our repeater pipets 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. Sometimes you can cross-reference with your tip box (e.g. I must be on the 4th column because I’ve used the first 3 columns worth of tips).
Another helper is color-coding. 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.
One of my favorite things, which I found a couple years ago, are “cluster tubes” which are kinda like giant PCR strip tubes – they’re strips of 8 tubes that fit in a deep well block frame and are great for doing larger-volume reactions without wasting a whole block. Plus, if you want, you can take out strips when you need to transfer liquid from them. I’ve found it’s easier to do this way because then you can see what you’re pipetting and make sure you’re not just pipetting up air bubbles.
Speaking of air bubbles… if you want to remove them without pipetting, try spinning them down in a centrifuge. Centrifuges often have swappable rotors so you can switch out the ones for tubes (of various sizes) for ones for plates. (yep – that’s what those tools next to the centrifuge are for!) A short pulse spin will draw the liquid down off the walls and below the air.
I was doing all this this morning when performing slot blot assays, which are a way to test protein-RNA binding. http://bit.ly/bindingaffinity ⠀
more on serial dilutions here: http://bit.ly/cerealdilutions
but the basic idea is you dilute something, then you dilute that something, and then you dilute that something, and then you dilute that something, and then…
each time you see “dilute” think “pipet”! My thumbs are not a fan, but my brain is because serial dilutions allow us to generate a series of exponentially-spaced (e.g. 1, 1/2, 1/4, 1/8, 1/16, 1/32…) samples covering a large concentration range. These dilutions are useful, but definitely not fun… they’re one of my very least parts of lab. So I definitely don’t want to have to repeat them! So here’s some tips I have.
Depending on the volumes you want it’s helpful to do them in strip tubes (like PCR strip tubes) or cluster tubes strips, deep well blocks, etc. – something where the liquid-holders are connected so you don’t mix them up. For today’s experiments I was doing 2-and-a-half blocks worth of cluster tube strips (with dilutions spanning 2 columns so I had 12 dilutions & 16 concentrations for each)
I can start by
- make the highest concentration at the concentration you want to start at then pipet the diluent (buffer (pH-stabilized salt water) in this case) into all the tubes except the first one which I save for the starting, highest concentration one that doesn’t need to get diluted. Then stick the starting one into the first tube and the 2nd tube.
- make a 2X initial concentration and then add that to diluent in the first well. so, in that case, you would add the same amount of diluent to all of the wells and then you add your sample to the first well and use that as your initial dilution. But, since you’re then diluting it, you need it to be 2X the desired starting concentration before you add it. For example, if you want to have 100uL of each concentration want to start at 400nM, either start by pipetting 100uL of 400nM sample into empty first well, or start by pipetting 100uL of 800nM sample into 100uL of buffer.
Usually what I do is I transfer then pipet up-down-up-down-up-down… (usually I do 5 times) ending on “up” then transfer to the next tube for the down⠀
When you’re making dilutions, you have to remember that you’re taking out part of your “V₂” each time – so if you do 10uL + 10uL = 20uL and then take 10uL of that for the next one you only have 10uL of that one left – not 20. So if you need 20, you should be doing 20 + 20 (actually if you need 20 you should be doing 25+25 or something because it’s always good to have a little extra to account for evaporation, tube/pipet sticking-to-ness, etc.) This is the same type of thing I mentioned earlier with maxing extra master mix.
Hope this random practical tip post can help make you a master of mixing and come to your rescue when you’ve got a lot of pipetting to do! 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!)
more on topics mentioned (& others) #365DaysOfScience All (with topics listed) 👉 http://bit.ly/2OllAB0⠀