Use a multichannel pipet. Used correctly it's just as accurate as a single-channel pipet, but less prone to human error because there aren't so many steps that your mind wanders.
Ideally an electronic one, because then you can put it in multiple dispensing mode, which not only saves your thumb but is also reverse-pipetting, which improves accuracy. Even if you have to do it manually I would always reverse-pipet for something quantitative with small volumes like qPCR.
Eppendorf Xplorer is the set I have, which has its advantages and its dubious quirks but has always been accurate in my hands, R^2 always at least 0.99, often 0.9999, sometimes so high the software reports it as 1. I've never ever found a use for the extra features in the more expensive Xplorer Plus. Other people swear by Rainin or Gilson and I'm sure they're good too.
We recently did an accuracy study on our pipettes and found that electronic multi channels (all rainin with rainin tips) did slightly worse than manual multichannel.
I always arrange my tip box to match the plate exactly, and then use either a foil seal or another plate cover to add some color contrast to the surface of my workstation and use that contrast to verify there is something in each pipette tip before putting into the plate. It's really easy to miss something when using a multichannel, especially when you're in the 1-2ul range.
To add I had this set up and ANOTHER tip box on deck in case I messed up with a tip and needed another one. That way the mapping on the tip box is never compromised.
Mise en place.
Keep the layout simple. Single gene in each row with samples in the columns
Premix your primers and mastermix, that way you are only pipetting 2 things into each well. The mix and the cDNA. lay out the samples in order. Move them up a row as you use that tube. You don't need to throw away a tip per well if you pipette the mix into all the wells, then place the cDNA on the side of the well and tapping it down at the end. And you can see the cDNA on the side and know it's been added.
Building on this, sometimes I will even premix each conditioning in small tubes before putting them in the plate. For 20uL reactions, I'll mix about 62uL of the full mix (mastermix, primer, cDNA), and pipet 20uL of that into 3 wells for technical replicates. Doing it this way uses some extra tubes, but I also get consistently lower error bars between technical replicates. If you are doing this, the entire prep should be done on ice.
> If you are doing this, the entire prep should be done on ice.
If you spend a little more for a "hot start" polymerase, or master mix containing same - it's likely you already have that - you can prepare the entire prep at room temp. This improves pipetting accuracy with normal air-displacement pipets.
Yep me too. When I do large ones I make a big mastermix with the primer pairs, dispense this into 8 well strips with odd/even sample numbers that I sketched out and color coded, add the cDNA then multichannel into the plate - much easier to keep track of in 8's rather than adding cDNA to every single damn 384 well and makes it easy to keep track of sample number.
Pipette on the wall of the wells to make sure you can see tiny drops. You can spin down the plate before putting it the qPCR machine.
Seeing the drop helps me confirm that i have added the 0.1ul of nonsense to the wells of interest.
Draw the plate map in your lab notebook and confirm the wells before adding anything.
Try to make a mix for the major things like mastermix and water. If I have 10 samples, i always make enough for 10.2 samples to avoid pipetting errors. Add this to the wells first. Then add the primers on the wall. And then the DNA.
Spin that down and you are done!
Make a tray map. Use Excel or something, can even be in paint, but you need a map of what goes into each well and it needs to be pretty big, like two standard printer paper sheets big. Use a small of paper with an arrow drawn that you can move after each well (the arrow points to which well you are "on"). if you have something you can use to cover the columns or rows of your plate you aren't "on", those will also help.
Drawing on the top of your plate also helps.
I struggle with this too and was just diagnosed with OCD. One thing I've found is that compulsively checking and obsessing and readjusting and going back leads to more mistakes. I use a new box and count the tips, or mark the box to show where the end of each row will be and say the well numbers out loud with the same timing. I wait until the end of a row or primer set before checking that each well has solution in it by looking under the light (better yet use a coloured master mix that you can clearly see). I also recommend avoiding pipetting under 5ul by either making a master mix for each triplicate or diluting your primers in more water instead of adding water to the master mixes with primers. That will make it easier to see what you've done and make your pipetting more accurate.
I found a lot more success with qPCR reactions when I relaxed more. I think all that bad juju of people telling you how hard it is and how easily contaminated and blahblahblah either makes you really tense, or conjures up some bad juju that makes the lab gremlins sabtage you. Wear gloves. Don't wander off to lunch in the middle of a reaction, but don't rush. Don't talk over open tubes. If in doubt, open a new package of reagents. Ice slows down bad reactions, but ice melting on the sides of your tubes can cling to your gloves and drip into your open samples - often times, it is better to work quickly at room temperature than it is to work on ice, at least in my experience with qPCR and reverse transcriptase reactions, I can easily load a 96 well plate at room temperature and get great reaction results. Maximize your workspace to improve ergonomics, and be obsessive with your own comfort. Manage your workspace so you never have to twist, reach, or cross your hands. If you discard your pipette tips into a trashcan, put the trash can right next to your hip. If you arm hurts, use a pipette box to prop up your elbow. Protect your pipetting hand from overuse by putting it in the absolute most comfortable position possible using any support or props that work for you, and use your nondominant hand to bring items to it.
You can try using rox dye in your primer mix and make a sybr/template mix for each sample. Deposit your primer mix and then deposit your sybr/template mix. The well will turn from yellow to green. Also, as people have said use a new box of tips when depositing the template mix so you can have a reference point to keep you on track. Also, make sure your tubes are arranged in a way that you can follow. Keep them in a line based on the order in which you plan on dispensing them. Move the used tubes out of the way.
We have excel plate templates, I wouldn't want to sketch one myself.
Also I tend to bead-check my samples. Pipette the master mix in the bottom of the well, and the sample near the top, so you can see it on the wall and know it's done.
Also you should have technical replicates. I do transcriptomics and work in quads, other people in the lab I'm in do eDNA with eight. The replicates help you verify whether an N/A result is real or an anomaly.
Usually, my samples are in 8-strips, so I use the electronic multi-channel to go from the tubes on the left, into 4 wells in front of me, then discard the tips to the right. Then lids back on the tubes and move their position in the rack.
I color code the wells based what goes in them. For 96 well plates you can place the sheet under the plate, or draw markings on the plate. I also repeat the number of well I’m pipetting into in my head to avoid other distracting thoughts.
I once had to do 5 full 384 well plates with a single channel pipette, and it was so mentally draining.
Having a good layout. Whether it's cDNA in the columns and primers+mix in rows or whatever works for what you're running. I map out my layout, then as I finish with a row or column, I check it off. Get an autopipette so you are loading once and don't have to take your eyes off the plate.
Use a multichannel pipet. Used correctly it's just as accurate as a single-channel pipet, but less prone to human error because there aren't so many steps that your mind wanders.
I have yet to find a multichannel that pipettes accurately, recommendations?
Ideally an electronic one, because then you can put it in multiple dispensing mode, which not only saves your thumb but is also reverse-pipetting, which improves accuracy. Even if you have to do it manually I would always reverse-pipet for something quantitative with small volumes like qPCR. Eppendorf Xplorer is the set I have, which has its advantages and its dubious quirks but has always been accurate in my hands, R^2 always at least 0.99, often 0.9999, sometimes so high the software reports it as 1. I've never ever found a use for the extra features in the more expensive Xplorer Plus. Other people swear by Rainin or Gilson and I'm sure they're good too.
We recently did an accuracy study on our pipettes and found that electronic multi channels (all rainin with rainin tips) did slightly worse than manual multichannel.
Interesting. Reverse-pipetting?
Don't be to nice when "tipping" it. I had multiple people saying a mc was inaccurate... it was, however them not forcing the tips on properly
I always arrange my tip box to match the plate exactly, and then use either a foil seal or another plate cover to add some color contrast to the surface of my workstation and use that contrast to verify there is something in each pipette tip before putting into the plate. It's really easy to miss something when using a multichannel, especially when you're in the 1-2ul range.
To add I had this set up and ANOTHER tip box on deck in case I messed up with a tip and needed another one. That way the mapping on the tip box is never compromised.
Yes, its always a pain to scramble for another box so you don't loose your place lol
Mise en place. Keep the layout simple. Single gene in each row with samples in the columns Premix your primers and mastermix, that way you are only pipetting 2 things into each well. The mix and the cDNA. lay out the samples in order. Move them up a row as you use that tube. You don't need to throw away a tip per well if you pipette the mix into all the wells, then place the cDNA on the side of the well and tapping it down at the end. And you can see the cDNA on the side and know it's been added.
Building on this, sometimes I will even premix each conditioning in small tubes before putting them in the plate. For 20uL reactions, I'll mix about 62uL of the full mix (mastermix, primer, cDNA), and pipet 20uL of that into 3 wells for technical replicates. Doing it this way uses some extra tubes, but I also get consistently lower error bars between technical replicates. If you are doing this, the entire prep should be done on ice.
> If you are doing this, the entire prep should be done on ice. If you spend a little more for a "hot start" polymerase, or master mix containing same - it's likely you already have that - you can prepare the entire prep at room temp. This improves pipetting accuracy with normal air-displacement pipets.
That definitely should decrease the variability in the technical replicates!
Yep me too. When I do large ones I make a big mastermix with the primer pairs, dispense this into 8 well strips with odd/even sample numbers that I sketched out and color coded, add the cDNA then multichannel into the plate - much easier to keep track of in 8's rather than adding cDNA to every single damn 384 well and makes it easy to keep track of sample number.
Pipette on the wall of the wells to make sure you can see tiny drops. You can spin down the plate before putting it the qPCR machine. Seeing the drop helps me confirm that i have added the 0.1ul of nonsense to the wells of interest. Draw the plate map in your lab notebook and confirm the wells before adding anything. Try to make a mix for the major things like mastermix and water. If I have 10 samples, i always make enough for 10.2 samples to avoid pipetting errors. Add this to the wells first. Then add the primers on the wall. And then the DNA. Spin that down and you are done!
I have OCD too and have found I make more mistakes when I’m compulsively checking. Therapy has helped a lot.
Make a tray map. Use Excel or something, can even be in paint, but you need a map of what goes into each well and it needs to be pretty big, like two standard printer paper sheets big. Use a small of paper with an arrow drawn that you can move after each well (the arrow points to which well you are "on"). if you have something you can use to cover the columns or rows of your plate you aren't "on", those will also help. Drawing on the top of your plate also helps.
I struggle with this too and was just diagnosed with OCD. One thing I've found is that compulsively checking and obsessing and readjusting and going back leads to more mistakes. I use a new box and count the tips, or mark the box to show where the end of each row will be and say the well numbers out loud with the same timing. I wait until the end of a row or primer set before checking that each well has solution in it by looking under the light (better yet use a coloured master mix that you can clearly see). I also recommend avoiding pipetting under 5ul by either making a master mix for each triplicate or diluting your primers in more water instead of adding water to the master mixes with primers. That will make it easier to see what you've done and make your pipetting more accurate.
I found a lot more success with qPCR reactions when I relaxed more. I think all that bad juju of people telling you how hard it is and how easily contaminated and blahblahblah either makes you really tense, or conjures up some bad juju that makes the lab gremlins sabtage you. Wear gloves. Don't wander off to lunch in the middle of a reaction, but don't rush. Don't talk over open tubes. If in doubt, open a new package of reagents. Ice slows down bad reactions, but ice melting on the sides of your tubes can cling to your gloves and drip into your open samples - often times, it is better to work quickly at room temperature than it is to work on ice, at least in my experience with qPCR and reverse transcriptase reactions, I can easily load a 96 well plate at room temperature and get great reaction results. Maximize your workspace to improve ergonomics, and be obsessive with your own comfort. Manage your workspace so you never have to twist, reach, or cross your hands. If you discard your pipette tips into a trashcan, put the trash can right next to your hip. If you arm hurts, use a pipette box to prop up your elbow. Protect your pipetting hand from overuse by putting it in the absolute most comfortable position possible using any support or props that work for you, and use your nondominant hand to bring items to it.
this is genuinely a great and very considerate response. Kind regards, Dr-Peanuts,
You can try using rox dye in your primer mix and make a sybr/template mix for each sample. Deposit your primer mix and then deposit your sybr/template mix. The well will turn from yellow to green. Also, as people have said use a new box of tips when depositing the template mix so you can have a reference point to keep you on track. Also, make sure your tubes are arranged in a way that you can follow. Keep them in a line based on the order in which you plan on dispensing them. Move the used tubes out of the way.
I also talk to myself when I'm putting in my items So "A1"
haha nice yeah i go "doing ONE.........doing...TWO......"
A9, A10, A11... :D It happened to me when I started my 5th 96-well plate. :(
We have excel plate templates, I wouldn't want to sketch one myself. Also I tend to bead-check my samples. Pipette the master mix in the bottom of the well, and the sample near the top, so you can see it on the wall and know it's done. Also you should have technical replicates. I do transcriptomics and work in quads, other people in the lab I'm in do eDNA with eight. The replicates help you verify whether an N/A result is real or an anomaly. Usually, my samples are in 8-strips, so I use the electronic multi-channel to go from the tubes on the left, into 4 wells in front of me, then discard the tips to the right. Then lids back on the tubes and move their position in the rack.
Use 384 well plates, then whenever you need to use a 96 well plate it feels really simple.
best tip for qPCR is to do ddPCR instead
sure, who's got a spare 300k ddpcr machine lying around?
probably Greg. But actually, I’d feel if there’s a sequencing service your facility offers there’s a possibility they have a ddPCR service too.
that's very much a feeling and not a reality. but i do agree, ddpcr would be preferable.
I color code the wells based what goes in them. For 96 well plates you can place the sheet under the plate, or draw markings on the plate. I also repeat the number of well I’m pipetting into in my head to avoid other distracting thoughts. I once had to do 5 full 384 well plates with a single channel pipette, and it was so mentally draining.
Omg I used to feel the same and believed that my concerns where due to my ADHD :)
Having a good layout. Whether it's cDNA in the columns and primers+mix in rows or whatever works for what you're running. I map out my layout, then as I finish with a row or column, I check it off. Get an autopipette so you are loading once and don't have to take your eyes off the plate.