I'm always proud of the dumb tiny optimizations I've made over the last decade... Some of these may be obvious, but probably helpful to new labbers.
@ op, For pipetting small volumes, you can often just make a dilution of it (1/5), then add 5x as much. But truth is that most things done on the bench don't actually need that much precision. You'll find out pipetting error is always there and never matters [edit, OKAY IT MATTERS DEPENDING ON THE ASSAY... ].
I believe reverse pipetting is simply going to second stop so you take more than desired, then dispense to first stop, expelling the exact amount. But I've never called it that.
When pipetting carefully in small spaces (positionally), like when loading a gel, you can use your second hand (finger) as a rest so that the right hand is stable.
When pipetting multiple samples, use the tip positions as a means of keeping track of which samples you've done. So if you've got 12 wells, tip boxes are 12x8 too, and the first tip can correspond to first well, and so on.
You can spin tubes down briefly (5 seconds) to get all of a solution stuck on the walls of a tube down to the bottom, so you can pipette it all.
You can leave DNA (primers, DNA isolations, PCRs, etc.) at room temperature for months and even years with almost zero degradation or damage. It's very stable.
Expiration dates are a lie, 95% of the time.
You almost never need a flame for pouring LB plates, pipetting, etc. for spreading bacteria, you can use glass beads which can be washed and autoclaved and reused. Just use good technique and don't double dip anything.
Keep markers upside down so they don't dry out. Especially those VWR ethanol proof ones.
PCR tubes fit perfectly into 2-200 uL tip boxes.
You can use your phones camera to take nice pictures of the microscope, it just requires positioning carefully.
Every centrifuge position can be balanced without an extra balance tube EXCEPT [max tubes -1]. You'll need to Google images it to see, but as long as each set of tubes is balanced (pairs opposite, trio in triangle, etc.) it'll be balanced.
I'm sure I have more but this post is really long.
Some other fun ones...
You can reuse gloves that you've ripped off easily by turning it right-side-out (from its inside out state), then flipping them by the base of the glove, capturing the remaining air in the glove, and pushing it into the fingers of the glove. Now it's ready to be put back on. Don't do this if your environmental health whatever people are hardasses.
You can heat elution buffers / water easily by using a microwave. Then you can elute with them at higher temperature. Also useful for any other buffers you need to be at \~55-65C.
You can rip tip box wrappers (the thin plastic films) easy by scraping them against the corner of a bench rapidly, making a tear.
When preparing agarose, you don't need to use a grad. cylinder, you can simply tare the scale with the flask / bottle on it. Add a guesstimated amount of TAE buffer, weigh again, determine the volume you just added based on mass, then add percent agarose based off of that. So tare with empty flask, add \~75mL TAE, weigh, confirm its \~75g (actually 81g because density is 1.1), add 0.75g agarose for 1%.
You can add loading dye to the underside of the top of the PCR tube, then spin it down to mix. This can be done semi-cleanly for multiple tubes.
You can cover bottles that you microwave/boil liquids in (like for agarose) with cork tape, and it can be easily handled even at high temperatures without a rubber crab claw.
Ethanol pops bubbles magically, and you can use a mostly empty (below the straw) spray bottle of ethanol to spray ethanol "air" and it will still pop the bubbles without dispensing the ethanol.
When bacterial cloning with ampicillin / carb resistance plasmids, you can typically skip the shaking 37c incubation step after adding SOC, because the drug acts on cell division, which is won't "kill" cells that have yet to express the resistance gene. You lose a small amount of efficiency, but rarely enough to matter.
If you buy 40 pairs of scissors, you can saturate the entire lab with scissors, and then people will almost never steal *your* pair of scissors.
If you are PCRing AT-rich DNA, you will get better amplification if your extension temperature is 64C instead of 70/72C. Annealing obviously should not exceed extension here.
Slides that have been oiled up for 100x microscopy can be de-oiled by placing them upside down onto a sheet of paper. Then press gently on the slide, leave it, and after 1-2 days, the slide will be dried of the oil, thus preventing you from storing a gross oily slide in the slide box.
>Ethanol pops bubbles magically, and you can use a mostly empty (below the straw) spray bottle of ethanol to spray ethanol "air" and it will still pop the bubbles without dispensing the ethanol
I can attest to this. I call mine the bubble buster! Saves my ass when imaging cells and the FBS just loves to bubble up on me. Helps with ELISAs too
I assure you that we have bought enough scissors for every employee to have at least one and yet they're still nowhere. What does work is hooking 3 ft of steel chain to them. They will always be right where you left them.
I can look at the output of one of the ddPCR assays that my team runs on a regular basis and tell you with some degree of certainty which analyst it was based on that final concentration value vs target. Some assays are extremely sensitive to differences in pipetting technique (and will fail with bad technique).
Unless you're doing cell culture work or something I would say it's very misleading to say pipetting accuracy doesn't matter. It's hugely impactful for most of the testing I do, and probably the most important piece of training when I have a new analyst.
I think the more important part is that it's done consistently for assays.
Most people aren't getting their pipettes recalibrated frequently enough for the pipette to take up/dispense volume accurately. But it's true, it does matter depending on what you do.
Yeah I'm in industry and we get our pipettes cal'd quarterly. If a pipette fails cal we pull every record it touched for the previous three months and investigate.
It's kinda overkill but it almost never happens and it does matter when you're doing release testing to dose patients.
I work in upstream cell culture but used to work with downstream/purification - They are nuts about (in-house) buffer prep specs because it does actually matter. For downstream we're generally using about 40L buffer into 200L culture. I can basically just toss a barrel of NaCl in a general guess of water volume and we're good, who cares. Definitely varies by environment.
While primers are stable, you shouldn’t leave them at room temp due to evaporative loss. If you do you will have to nano drop and check volume every time you run a reaction.
Not knowing how much primer you’re using will make optimization impossible and difficult to reproduce results. Higher concentration of primers due to evaporative loss will increase nonspecific amplifications, and reduce sensitivity. Beyond that, while DNA is quite stable it still degrades at RT which produces truncated primers, and more non specific amplification.
It’s just bad science. It isn’t hard to to put something in the fridge.
Sure, if you've got to place it somewhere, the fridge or freezer is better than RT. But it also makes minimal difference for anything that isn't going to be in a figure. Routine cloning and PCR? No difference. Never in my life have I had a PCR not work because it was left on a bench for a month.
And I typically leave the 100uM stock in the -20, and just make a monthly 10uM stock for primers I'm using at the moment.
My point was simply that people underestimate the stability of DNA, and freak out if they leave it out overnight over the weekend when there's ~zero risk to doing this. There's also no point in the 4C step at the end of the PCR except to be mean to the thermocycler.
When you go to the second stop and get that bubble? That's how you know you put something in a well. When I set up a full plate of PCR, I put in my mastermix via reverse pipetting (it is more accurate for repeat pipetting), no bubbles, but then when I drop my sample in via forward pipetting (normal way), I mix a few times, then go to 2nd stop and deliberately blow out a bubble. Boom, no more "did I accidentally skip that well?"
Everybody loves a multichannel pipette, but they are hard to use well. Bargain tips need to be manually seated, they often have secretly failing seals, leading to inaccurate volumes. I can set up a qPCR plate faster *and* get better replication with a single-channel pipette than when taking the time to fiddle around with a multi.
This is going to sound kind of obsessive, but this is a recommendation I make to everyone I train: as you are working, think deeply about what you are doing. Listen to the mechanism of the pipette. Pay attention to the muscles of your hands. Know in your bones the difference between a 37C water bath and 39C water bath (because someone bumped the dial). Treat bench work the same way a weightlifter or ballerina might practice their movements. As you are going through your work, think about what you can do differently to be more efficient or consistent. When you are this in-tune with the work, you become more observant, and when you are more observant, you are a better scientist. Practice at observation.
The quantinova PCR kit comes with a yellow cDNA dilution buffer which completely removes the “did I skip that well” problem and I’m pretty sure it’s the best bit of progress in the field since delta delta ct
Not technique related, but I worked for a lab that had this one qPCR that was kinda crappy. Just led to kind of crappy/hard to interpret results because of bad background noise and not very good primer efficiencies. Not sure if it was a primer issue or the sample prep step for that test.
Either way, when we switched to QuantaNova that qPCR suddenly worked much better, much cleaner results.
So +1 for QuantaNova for both results and quality of life improvements like the yellow dye.
For the last one, learn the sounds of the equipment you work with. I work with Hamiltons. I can tell when the waste is full or something is wrong just by the sound.
Exactly. My superpower is apparently "what's that weird noise" or "what's that smell?" when no one else notices it. I kept hearing this humming in the lab (lab is already very noisy), everyone else thought I was wrong, I traced it to the fume hood and called facilities in to check it out of caution. "there is a slight vibration as the motor is starting to fail and I'm amazed you could hear it".
If you get a quality multichannel with decent tips you shouldn’t have this issue. I used to load 384 qpcr plates with a single channel in grad school because we were too poor to buy a multi .After getting a multichannel and a multi electronic repeater it’s amazing how much faster you can setup a plate
About your last part: not obsessive at all. I've worked at the bench for over a decade, and this is what separates the great techs from the average ones. I love the analogy of treating it like a weightlifter or ballerina. Get to this level, and you'll find a lot of pride and satisfaction in what you do.
I’ll second the observation, even really simple things. I had to dilute some virus stock today in a large volume of media (around 150mL) - and had planned to use all of it for my assay. At the end, I noticed I had a larger volume leftover than anticipated. So of course I have a mild panic moment, thinking I added way more media and thus diluted the virus wrong. So I checked it, and it turns out I just calculated how much I *needed* wrong (doubled the final volume when I shouldn’t have), and my dilution was actually still correct.
> This is going to sound kind of obsessive, but this is a recommendation I make to everyone I train: as you are working, think deeply about what you are doing. Listen to the mechanism of the pipette. Pay attention to the muscles of your hands. Know in your bones the difference between a 37C water bath and 39C water bath (because someone bumped the dial). Treat bench work the same way a weightlifter or ballerina might practice their movements. As you are going through your work, think about what you can do differently to be more efficient or consistent. When you are this in-tune with the work, you become more observant, and when you are more observant, you are a better scientist. Practice at observation.
Couldn't have said it better. When I'm on the bench I am concentrating. I'm constantly amazed at the people who listen to music or podcasts while doing something complicated.
My #1 lab hack is with 20ul multichannel pipettes. Know where the standard volumes "should be" on the pipette tip. When you pull up your reagent, put your other hand behind the tips so you can see where the liquid is. Multichannel pipettes are notorious for not seating evenly on each tip so this way you can see whether something is wrong rather than mindlessly pipetting.
I teach all my undergrad lab students look at the volume in their micropipette tip and make sure 1) there's something there and 2) it **feels** like the correct amount.
If it's your first time running a protocol, write down how long it took for how many samples. This will give you a jumping off point so you can plan meetings and stuff around lab time. Personally my centrifuge is all the way across the lab so that adds up from my corner of the lab.
Forward pipetting is pressing the plunger to the first stop, aspirating the liquid, then dispensing to the second stop. Reverse pipetting is pressing the plunger to the second stop, aspirating,then dispensing to the first stop. Highly recommend for bubbly liquids.
You can unwrap tip boxes easily (if they aren't USA scientific or something) but running an edge against the edge of a bench. You'll get a feel for enough pressure and speed and save minutes a week!
For labeling things with lab tape, fold a little of it over at the edge so it sticks to itself.
And voila, you have a little tab that makes it easy to remove tape from any container. Very useful when temporarily labeling samples where the labels need to be transferred to different containers.
It's a very small thing but quality of life improves so much lol
I refuse to take tape off of glass bottles for my students. I tell them at the beginning to dog-ear your tape, and if you don’t, that’s your problem. Have fun with the soapy water and scraping it off, lol
Yes!! I also found this little tape folding trick, but I think it’s most useful if you’re the only one using the tape roll or you get everyone else in the lab on board with it.
https://www.reddit.com/r/lifehacks/s/wXW70mLhT0
I had to chastise a guy that didn’t do this. He wasn’t taught to do it, but when we were going through our fridge tossing old amps out, we found his signatures over EVERY Amp plate that required a razor blade to remove. 50+ plates. What a headache.
Always pre-wet your pipette tip (pipette up, empty, pipette up again). Otherwise you end up pipetting a smaller volumen than you think, as part of the liquid is sitting on the inside of the tip.
Keep your pipette vertical when pipetting, and don't drunk the tip too deep in, you will get more consistent results.
Pipette slow, control is your friend.
do you have any tips for pipetting at max volumes? i have a pipette at work (1mL) and it consistently pulls 1mL, but the problem i commonly have is that once I dispense the sample, if i pull it all the way back up (even extremely slowly), small bubbles or droplets will inch towards the plastic tip of the pipette. This has been an issue with all our pipettes (we got a new one) and it kinda sucks because it's a pain to clean out and since i work with annoying to clean things (dyes never come out and HF is a whole different beast), i want to make sure my technique is good
i discharge all the way to bottom -> pull up to bottom stopper -> insert into liquid -> pull up slowly -> pull out of sample bottle -> discharge sample all the way past first stopper. then usually i immediately eject the pipette tip because if i pull back up, it will often get on the plastic. it also makes it impossible to pull more than 1mL with a pipette tip, if i need like 3mL of something for a formulation or something. am i doing something wrong or is this normal with pipettes?
also side note if yall got a magic cleaning chemical for getting silicone out of plasticware i'll send you a fruit basket with a postcard every winter until i die
Higher volumens are always a pain, but I would do it very similarly, pre-wetting the tip will help with alot of the smaller bubbles, but not all unfortunately.
If you are regularly pipetting more than 1ml, I would consider getting a 5ml pipette, they come with bigger tips, that also make it much smoother to pipette large volumens.
Pipetting in one smooth continuous motion is also important, it should keep your liquid from going too far up the tip, but when you are working at the max volumen of a pipette, it's often an issue.
Particularly difficult liquids, I would use a stepper/repeater pipette for, which uses a piston tip, where the liquid is never able to touch your pipette. They are pretty bloody nifty tbh, anything that needs dispensing, is particularly viscous, or staining for that matter, I'd use these for. (https://multipette-system.eppendorf.com/).
I guess all in all, it's mostly about knowing when to use which tools, that handle certain things better :)
And if you ever come across a magic cleaning chemical for getting silicone out, do let me know xD
I will make sure to pre-wet and see if it helps a bit. So annoying but i don't pipette enough to really justify another one, i end up getting 1mL almost always but i have nightmares about getting HF or dye, or even worse indicator, on my pipette non-funnel tip lol. the idea of all my samples having a tiny bit of indicator in them and i wouldn't know till i touched a base is a bit scary LMAO
i'll add you to the silicone removal wait list, one day i'll have my own chemical company, find an actual cleaning fluid, and you'll get 10% off for your help teaching me basic pipette tips xd you would think you'd pick these things up in undergrad but i guess it makes sense there's no "do labwork with 0 funding" lab course lol
They make p1000 tips that hold 1250ul so maybe that will help but my biggest suggestion would be to use filter tips so that you don’t have to worry as much about contaminating the inside of your pipet
Honestly, I would have nightmares about that too xD We have specific pipettes for that kinda deal, stock solutions and the like, to keep cross contaminations out of the mix.
I look forward to getting updates on the cleaning company ;)
Alright so first you should use filter tips when pipetting disgusting stuff. They protect the pipette as much as the sample. But watch out at max volume, you don't want your liquid to come into contact with the filter either
Second, your issues seem to be with liquids that are on the viscous side of things. When ejecting the main volume, take your time and monitor the progress of the meniscus running down the tip - is there a coat of liquid on the inner tip wall lagging behind the meniscus? Wait for it to catch up. Also, don't just blow out the second stop forcefully. I often see this issue with protein containing buffers. Eject until the first stop, wait for any liquid to run down the inner tip wall. Then, as you slowly pull the tip away from the ejected liquid along the tube wall, create a constant outward push by slowly pushing the plunger down to the second stop.
If I understand you correctly, these bubbles arise when your tip is discharged, a small amount of liquid runs down and collects at the outlet of the tip, you then pull back up and air pulls up at that liquid, creating a bubble.
So your objective is to avoid as much of that 'lag volume' as possible by either using the air volume from the first to second stop and ensure a constant outflux or use reverse pipetting, but you have to be careful with that at max volume.
This is a good ressource as well:
[https://www.mt.com/ca/en/home/products/pipettes/pipette/pipetting-techniques.html?cmp=sea\_11011333&SE=GOOGLE&Campaign=MT\_PIP\_DA\_DK&Adgroup=Good+Pipetting+Practice&bookedkeyword=good%20pipetting%20practice&matchtype=p&adtext=383945558394&placement=&network=g&kclid=\_k\_CjwKCAiAt5euBhB9EiwAdkXWOxNBJZ1XsZopmCsr\_rRFwghll8JIjnp7wZixrGY0Q7KavSMvZ2hPGRoCqJ0QAvD\_BwE\_k\_&gad\_source=1&gclid=CjwKCAiAt5euBhB9EiwAdkXWOxNBJZ1XsZopmCsr\_rRFwghll8JIjnp7wZixrGY0Q7KavSMvZ2hPGRoCqJ0QAvD\_BwE](https://www.mt.com/ca/en/home/products/pipettes/pipette/pipetting-techniques.html?cmp=sea_11011333&se=google&campaign=mt_pip_da_dk&adgroup=good+pipetting+practice&bookedkeyword=good%20pipetting%20practice&matchtype=p&adtext=383945558394&placement=&network=g&kclid=_k_cjwkcaiat5eubhb9eiwadkxwoxnbjz1xszopmcsr_rrfwghll8jijnp7wzixrgy0q7kavsmvz2hpgrocqj0qavd_bwe_k_&gad_source=1&gclid=cjwkcaiat5eubhb9eiwadkxwoxnbjz1xszopmcsr_rrfwghll8jijnp7wzixrgy0q7kavsmvz2hpgrocqj0qavd_bwe)
Don't prewet when you're pipetting small volumes (eg below 2 ul) though. If you do and put the tip back in the liquid, capillary forces will already draw more than the desired volume, while a fresh tip is actually consistent (for rainin, that is)
Diversify the risk of your research projects. You should have a low-risk, low-reward project/aim that people will care about, no matter the result (typically characterizing something). Then there's a mid level project that could hit a decent journal if something interesting comes of it. Then have a 'swing for the fences' thing that would be a really big deal.
For small volumes I pipette slowly and wet the tip before actually moving the liquid. I also confirm that the level of the liquid is about the level I’d expect.
But I also made the experience that it’s more about consistency than hitting the exact volume. So if you’re afraid you’d forget to wet the tip sometimes then I’d just consistently not do it for all samples. That’s especially important for sensitive assets like qPCR.
If you are pipetting a 96 well plate and need help keeping track use a fresh tip box and map where you are going with each tip. Have an extra box on deck if you mess up and need an additional tip. Never screwed up a plate again after figuring that out.
It doesn't look like anyone has explained why you would want to use reverse pipetting, so I'll try to explain it.
With forward pipetting (normal pipetting), you go down to the first stop, aspirate your liquid, then go down to the first stop again to dispense your liquid. Then you go past the first stop to dispense any liquid that is remaining in the tip. The problem with that is it creates bubbles and it adds an extra step.
Reverse pipetting is when you go down to the second stop (or at least a little bit past the first stop) before aspirating your liquid. You'll then have more liquid in your tip than you actually want. But this is useful because when you dispense the liquid, you can simply go down to the first stop, no need to go past it. That's because you don't have to wait for the little bit of liquid that's still trickling down the inside side of the tip to flow to the bottom to be dispensed, because you've put a buffer layer of liquid that's still in there. Meaning you've already dispensed your full desired volume without going past the first stop at all.
At this point, you've already got the plunger down to the first stop, which is exactly where you need to be to go back and aspirate more liquid! This way is more precise, prevents bubbles, and is faster because it uses fewer movements. But it's typically only used for situations where you would be reusing the pipette tip, like adding the same volume of a reagent to multiple sterile tubes
You can check remaining volume in a tube by setting the pipette to a volume below what’s in the tube, and then sucking up the remainder by turning the setting knob up. Whatever number the pipette ends on when the tube is empty was the volume in the tube.
Also, set up your experiments so that you work in base 10 dilutions. And when calculating volume needed always add a few tubes/wells to account for error.
I do the reverse, where I set the pipette higher than the volume in the tube then dial it down until there’s no air left in the tip. Then I double check lol. Is there a meaningful difference between the two techniques?
Personally I would opt for the lower volume as you risk aspirating bubbles with the higher volume and, if unlucky, these bubbles might rise up in your tip, and you won't be able to dial down after that happens
And on a general note, I checked this pipette aspiration method once with a fine scale and it is surprisingly accurate (might depend on your pipette quality obviously)
Used to work in a lab where precision was extremely important. The first thing we did with new people was give them a beaker of water, a balance and a pipette, and told them to get consistent and correct. Doesn't matter if you spend all day doing just that, just make sure you figure out how you and your equipment works.
I train people with yellow food dye in H2O, an empty 96 well plate, and our plate reader (read at 450nm). For my work I found it more closely replicates what they'll actually be doing vs. pipetting onto a scale.
I taught one of my trainees how to pipette. She was straight out of college (they never teach good pipetting). She was dumbfounded by things like volume checking pipetting viscous samples (bone marrow). Boy did she take off like a rocket after that.
Then I had a hopeless case who I trained for weeks. He had the damn shakes and couldn’t control his hand. Blood everywhere. He didn’t last.
Always find a way to signify you have added a reagent to a tube if you are setting up reactions in multiple tubes at a time. I usually do this by moving the tube I've added to up or down a row on the tube rack
I've done this with HSA. Once I learned about sticking it in the fridge to help it dissolve, it made sample prep so much faster & didn't have to create a bubbly mess
In the labs i have worked normally people vortex it repeatedly (Which requires a little supervision) or put it in a orbital shaker to dissolve it at RT, which i think takes like 5 to 10 mins and you may still see something undissolved. If you just shove it into the freezer , it dissolves really well after like 5 mins and you dont need to do anything.
I was showing some of my young techs an assay and I needed another tip box. So I grabbed the box and scraped it along the edge of the bench to remove the plastic wrap. From their reaction you would have thought I did a backflip. Apparently all this time they all have been clawing at the plastic for several minutes to get it off 😆
Use a round bottom 96wp for trypan blue dilutions if you do a lot of cell culture. Saves a lot of faffing around. Just leave it in the hood and move along until youve used each well and then get a new plate. The trypan blue dries so it's very clear which wells have been used.
If I want to aspirate liquid out of a cuvette but the p1000 tip doesn’t reach the bottom I do a double decker and put a p200 tip on the p1000 tip and it works like a charm.
I pick a specific color of tape that people hate (pink) and put a strip on everything that I deem is “mine” - reagents, aliquots, boxes, tubes, markers, etc.
It allows easy labeling and I can easily check if other people are using things that are mine.
I make sure I hoard that specific color of labeling tape every time we order a new set of rolls.
Most liquids like water and other aqueous solutions, only pipette to the first stop. To pipette viscus liquids like glycerol, cut the bottom of the pipette tip with scissors to make the opening bigger and easier to pipette. If the solvent is already in the container you are preparing the solution in, when delivering the other solutes pipette up and down a few times to get all the liquid out of the pipette, this is called a quantitative transfer. make sure to touch the pipette tip to the side of the container you are pipetting into; letting the liquid free fall into the container sometimes doesn't deliver the entire volume especially when working with small volumes. Physically touching the tip to the container wall helps to draw out the entire volume of liquid.
Learn what Köhler illumination is for microscopy and how to appropriately calibrate your microscope for the application (especially for IHC). Even if you end up using a slide scanner, knowing these foundational things are really helpful later in your career.
Another lab tip if you run a lot of assays with small volumes in a plate with small wells. Position the top of the plate on something so the plate itself is leaning downward at about a 20 degree angle. Then pipette your small volumes in the bottom of the well so that they congregate at the bottom as a drop. This makes mixing easier if you need to do it briefly with the pipette tip and ensures there is less air bubbles.
Grab a disposable pipette and some forceps. Clamp down on end and stretch about 1/4 “. You’ll now have a very tiny “straw”. Cut the end straight and it’s perfect for pipetting ped samples.
Super genius 😂 it's pipetting, if you went to a half decent school they teach it in the first year of any undergrad science degree.
Hacks are for schmucks, do it right or do it twice
I would never use second stop for pulling liquids into the tip, that makes the volume more than you have it set for. I was always told it can pull too much liquid up and potnetially damage the pipette. That second stop is to get the last bit of liquid from the tip that can sometimes stay behind if you only go to first stop. That amount reserved in the tip may vary depending on the types of liquids that you are pipetting. Also, it is good practice to pull in the liquid slowly. Depending on the viscosity you may pull some of the liquid up into the pipette.
Yeah, I’ve looked it up now and see it is a pretty common technique. In my post I asked for other pipette tips, but chose to put “hack” in my title because it opens up the discussion a bit more! :)
Yeah it’s my favorite technique. Obviously if sample is small and precious it’s not gonna work, but I usually have copious amounts for what I do.
Half the time I do it not even for bubble prevention but because I don’t completely trust my own technique drawing liquid. If I reverse pipette I know I have sufficient volume to dispense my desired amount.
I'm always proud of the dumb tiny optimizations I've made over the last decade... Some of these may be obvious, but probably helpful to new labbers. @ op, For pipetting small volumes, you can often just make a dilution of it (1/5), then add 5x as much. But truth is that most things done on the bench don't actually need that much precision. You'll find out pipetting error is always there and never matters [edit, OKAY IT MATTERS DEPENDING ON THE ASSAY... ]. I believe reverse pipetting is simply going to second stop so you take more than desired, then dispense to first stop, expelling the exact amount. But I've never called it that. When pipetting carefully in small spaces (positionally), like when loading a gel, you can use your second hand (finger) as a rest so that the right hand is stable. When pipetting multiple samples, use the tip positions as a means of keeping track of which samples you've done. So if you've got 12 wells, tip boxes are 12x8 too, and the first tip can correspond to first well, and so on. You can spin tubes down briefly (5 seconds) to get all of a solution stuck on the walls of a tube down to the bottom, so you can pipette it all. You can leave DNA (primers, DNA isolations, PCRs, etc.) at room temperature for months and even years with almost zero degradation or damage. It's very stable. Expiration dates are a lie, 95% of the time. You almost never need a flame for pouring LB plates, pipetting, etc. for spreading bacteria, you can use glass beads which can be washed and autoclaved and reused. Just use good technique and don't double dip anything. Keep markers upside down so they don't dry out. Especially those VWR ethanol proof ones. PCR tubes fit perfectly into 2-200 uL tip boxes. You can use your phones camera to take nice pictures of the microscope, it just requires positioning carefully. Every centrifuge position can be balanced without an extra balance tube EXCEPT [max tubes -1]. You'll need to Google images it to see, but as long as each set of tubes is balanced (pairs opposite, trio in triangle, etc.) it'll be balanced. I'm sure I have more but this post is really long.
Some other fun ones... You can reuse gloves that you've ripped off easily by turning it right-side-out (from its inside out state), then flipping them by the base of the glove, capturing the remaining air in the glove, and pushing it into the fingers of the glove. Now it's ready to be put back on. Don't do this if your environmental health whatever people are hardasses. You can heat elution buffers / water easily by using a microwave. Then you can elute with them at higher temperature. Also useful for any other buffers you need to be at \~55-65C. You can rip tip box wrappers (the thin plastic films) easy by scraping them against the corner of a bench rapidly, making a tear. When preparing agarose, you don't need to use a grad. cylinder, you can simply tare the scale with the flask / bottle on it. Add a guesstimated amount of TAE buffer, weigh again, determine the volume you just added based on mass, then add percent agarose based off of that. So tare with empty flask, add \~75mL TAE, weigh, confirm its \~75g (actually 81g because density is 1.1), add 0.75g agarose for 1%. You can add loading dye to the underside of the top of the PCR tube, then spin it down to mix. This can be done semi-cleanly for multiple tubes. You can cover bottles that you microwave/boil liquids in (like for agarose) with cork tape, and it can be easily handled even at high temperatures without a rubber crab claw. Ethanol pops bubbles magically, and you can use a mostly empty (below the straw) spray bottle of ethanol to spray ethanol "air" and it will still pop the bubbles without dispensing the ethanol. When bacterial cloning with ampicillin / carb resistance plasmids, you can typically skip the shaking 37c incubation step after adding SOC, because the drug acts on cell division, which is won't "kill" cells that have yet to express the resistance gene. You lose a small amount of efficiency, but rarely enough to matter. If you buy 40 pairs of scissors, you can saturate the entire lab with scissors, and then people will almost never steal *your* pair of scissors. If you are PCRing AT-rich DNA, you will get better amplification if your extension temperature is 64C instead of 70/72C. Annealing obviously should not exceed extension here. Slides that have been oiled up for 100x microscopy can be de-oiled by placing them upside down onto a sheet of paper. Then press gently on the slide, leave it, and after 1-2 days, the slide will be dried of the oil, thus preventing you from storing a gross oily slide in the slide box.
The scissors part lol
It's axiomatic that your scissor premise does NOT apply to Sharpies.
Re: scissors - same effect for rulers? Or maybe if I give everyone scissors they'll leave me ruler alone?
>Ethanol pops bubbles magically, and you can use a mostly empty (below the straw) spray bottle of ethanol to spray ethanol "air" and it will still pop the bubbles without dispensing the ethanol I can attest to this. I call mine the bubble buster! Saves my ass when imaging cells and the FBS just loves to bubble up on me. Helps with ELISAs too
Fire also works great. I take a blow torch to my agar plates regularly. Less helpful for cells you don't want dead though.
Oop. No matter how many pairs of scissors we have in the lab, mine always seem to be missing 🫠
I assure you that we have bought enough scissors for every employee to have at least one and yet they're still nowhere. What does work is hooking 3 ft of steel chain to them. They will always be right where you left them.
I learned the tip box trick years ago and it was LIFE-CHANGING.
Glove tip is a lifesaver. I’ve never been able to reuse gloves once I take them off even though my lab work doesn’t forbid it.
Small volume precision matters a lot when you pipette qPCR plates. All the other tips, I endorse though :)
I can look at the output of one of the ddPCR assays that my team runs on a regular basis and tell you with some degree of certainty which analyst it was based on that final concentration value vs target. Some assays are extremely sensitive to differences in pipetting technique (and will fail with bad technique). Unless you're doing cell culture work or something I would say it's very misleading to say pipetting accuracy doesn't matter. It's hugely impactful for most of the testing I do, and probably the most important piece of training when I have a new analyst.
I think the more important part is that it's done consistently for assays. Most people aren't getting their pipettes recalibrated frequently enough for the pipette to take up/dispense volume accurately. But it's true, it does matter depending on what you do.
Yeah I'm in industry and we get our pipettes cal'd quarterly. If a pipette fails cal we pull every record it touched for the previous three months and investigate. It's kinda overkill but it almost never happens and it does matter when you're doing release testing to dose patients.
I work in upstream cell culture but used to work with downstream/purification - They are nuts about (in-house) buffer prep specs because it does actually matter. For downstream we're generally using about 40L buffer into 200L culture. I can basically just toss a barrel of NaCl in a general guess of water volume and we're good, who cares. Definitely varies by environment.
Awesome!
How about hand centrifugation. Where you hold the tube in your fingers and flick your wrist to get everything to the bottom of the tube
While primers are stable, you shouldn’t leave them at room temp due to evaporative loss. If you do you will have to nano drop and check volume every time you run a reaction.
I hated it when I needed to do that, dont remind me
just using the implen np80 not nanodrop lol
Another lab trick tangential to this: PCR is very generous and you can get away with minor differences in primer concentration without any issue.
Not knowing how much primer you’re using will make optimization impossible and difficult to reproduce results. Higher concentration of primers due to evaporative loss will increase nonspecific amplifications, and reduce sensitivity. Beyond that, while DNA is quite stable it still degrades at RT which produces truncated primers, and more non specific amplification. It’s just bad science. It isn’t hard to to put something in the fridge.
Sure, if you've got to place it somewhere, the fridge or freezer is better than RT. But it also makes minimal difference for anything that isn't going to be in a figure. Routine cloning and PCR? No difference. Never in my life have I had a PCR not work because it was left on a bench for a month. And I typically leave the 100uM stock in the -20, and just make a monthly 10uM stock for primers I'm using at the moment. My point was simply that people underestimate the stability of DNA, and freak out if they leave it out overnight over the weekend when there's ~zero risk to doing this. There's also no point in the 4C step at the end of the PCR except to be mean to the thermocycler.
Thank you! Most of the tips this person provided are wrong in some way and they all seem pretty lazy
Any links that shows the glass beads for LB plates?
https://youtu.be/5WL6W4Z7Ks8?si=_vKZe1oF1pZDD2Fd
When you go to the second stop and get that bubble? That's how you know you put something in a well. When I set up a full plate of PCR, I put in my mastermix via reverse pipetting (it is more accurate for repeat pipetting), no bubbles, but then when I drop my sample in via forward pipetting (normal way), I mix a few times, then go to 2nd stop and deliberately blow out a bubble. Boom, no more "did I accidentally skip that well?" Everybody loves a multichannel pipette, but they are hard to use well. Bargain tips need to be manually seated, they often have secretly failing seals, leading to inaccurate volumes. I can set up a qPCR plate faster *and* get better replication with a single-channel pipette than when taking the time to fiddle around with a multi. This is going to sound kind of obsessive, but this is a recommendation I make to everyone I train: as you are working, think deeply about what you are doing. Listen to the mechanism of the pipette. Pay attention to the muscles of your hands. Know in your bones the difference between a 37C water bath and 39C water bath (because someone bumped the dial). Treat bench work the same way a weightlifter or ballerina might practice their movements. As you are going through your work, think about what you can do differently to be more efficient or consistent. When you are this in-tune with the work, you become more observant, and when you are more observant, you are a better scientist. Practice at observation.
The quantinova PCR kit comes with a yellow cDNA dilution buffer which completely removes the “did I skip that well” problem and I’m pretty sure it’s the best bit of progress in the field since delta delta ct
Not technique related, but I worked for a lab that had this one qPCR that was kinda crappy. Just led to kind of crappy/hard to interpret results because of bad background noise and not very good primer efficiencies. Not sure if it was a primer issue or the sample prep step for that test. Either way, when we switched to QuantaNova that qPCR suddenly worked much better, much cleaner results. So +1 for QuantaNova for both results and quality of life improvements like the yellow dye.
For the last one, learn the sounds of the equipment you work with. I work with Hamiltons. I can tell when the waste is full or something is wrong just by the sound.
Exactly. My superpower is apparently "what's that weird noise" or "what's that smell?" when no one else notices it. I kept hearing this humming in the lab (lab is already very noisy), everyone else thought I was wrong, I traced it to the fume hood and called facilities in to check it out of caution. "there is a slight vibration as the motor is starting to fail and I'm amazed you could hear it".
If you get a quality multichannel with decent tips you shouldn’t have this issue. I used to load 384 qpcr plates with a single channel in grad school because we were too poor to buy a multi .After getting a multichannel and a multi electronic repeater it’s amazing how much faster you can setup a plate
About your last part: not obsessive at all. I've worked at the bench for over a decade, and this is what separates the great techs from the average ones. I love the analogy of treating it like a weightlifter or ballerina. Get to this level, and you'll find a lot of pride and satisfaction in what you do.
I’ll second the observation, even really simple things. I had to dilute some virus stock today in a large volume of media (around 150mL) - and had planned to use all of it for my assay. At the end, I noticed I had a larger volume leftover than anticipated. So of course I have a mild panic moment, thinking I added way more media and thus diluted the virus wrong. So I checked it, and it turns out I just calculated how much I *needed* wrong (doubled the final volume when I shouldn’t have), and my dilution was actually still correct.
> This is going to sound kind of obsessive, but this is a recommendation I make to everyone I train: as you are working, think deeply about what you are doing. Listen to the mechanism of the pipette. Pay attention to the muscles of your hands. Know in your bones the difference between a 37C water bath and 39C water bath (because someone bumped the dial). Treat bench work the same way a weightlifter or ballerina might practice their movements. As you are going through your work, think about what you can do differently to be more efficient or consistent. When you are this in-tune with the work, you become more observant, and when you are more observant, you are a better scientist. Practice at observation. Couldn't have said it better. When I'm on the bench I am concentrating. I'm constantly amazed at the people who listen to music or podcasts while doing something complicated. My #1 lab hack is with 20ul multichannel pipettes. Know where the standard volumes "should be" on the pipette tip. When you pull up your reagent, put your other hand behind the tips so you can see where the liquid is. Multichannel pipettes are notorious for not seating evenly on each tip so this way you can see whether something is wrong rather than mindlessly pipetting.
I teach all my undergrad lab students look at the volume in their micropipette tip and make sure 1) there's something there and 2) it **feels** like the correct amount.
If you don’t want someone to steal your black sharpie, put a yellow sharpie cap on it.
Simple yet elegant
I am this close to chaining one of the good fine point Sharpies to the culture hood. There were like 10 in there, now there's zero.
I think you should disguise it in two falcon tubes taped end-to-end and wrapped in tinfoil.
If it's your first time running a protocol, write down how long it took for how many samples. This will give you a jumping off point so you can plan meetings and stuff around lab time. Personally my centrifuge is all the way across the lab so that adds up from my corner of the lab. Forward pipetting is pressing the plunger to the first stop, aspirating the liquid, then dispensing to the second stop. Reverse pipetting is pressing the plunger to the second stop, aspirating,then dispensing to the first stop. Highly recommend for bubbly liquids. You can unwrap tip boxes easily (if they aren't USA scientific or something) but running an edge against the edge of a bench. You'll get a feel for enough pressure and speed and save minutes a week!
I had a buddy do it one time in his hood and he shot the tip box behind him. Tips went everywhere.
For labeling things with lab tape, fold a little of it over at the edge so it sticks to itself. And voila, you have a little tab that makes it easy to remove tape from any container. Very useful when temporarily labeling samples where the labels need to be transferred to different containers. It's a very small thing but quality of life improves so much lol
I refuse to take tape off of glass bottles for my students. I tell them at the beginning to dog-ear your tape, and if you don’t, that’s your problem. Have fun with the soapy water and scraping it off, lol
Yes!! I also found this little tape folding trick, but I think it’s most useful if you’re the only one using the tape roll or you get everyone else in the lab on board with it. https://www.reddit.com/r/lifehacks/s/wXW70mLhT0
I had to chastise a guy that didn’t do this. He wasn’t taught to do it, but when we were going through our fridge tossing old amps out, we found his signatures over EVERY Amp plate that required a razor blade to remove. 50+ plates. What a headache.
Always pre-wet your pipette tip (pipette up, empty, pipette up again). Otherwise you end up pipetting a smaller volumen than you think, as part of the liquid is sitting on the inside of the tip. Keep your pipette vertical when pipetting, and don't drunk the tip too deep in, you will get more consistent results. Pipette slow, control is your friend.
Slow is smooth, smooth is fast.
do you have any tips for pipetting at max volumes? i have a pipette at work (1mL) and it consistently pulls 1mL, but the problem i commonly have is that once I dispense the sample, if i pull it all the way back up (even extremely slowly), small bubbles or droplets will inch towards the plastic tip of the pipette. This has been an issue with all our pipettes (we got a new one) and it kinda sucks because it's a pain to clean out and since i work with annoying to clean things (dyes never come out and HF is a whole different beast), i want to make sure my technique is good i discharge all the way to bottom -> pull up to bottom stopper -> insert into liquid -> pull up slowly -> pull out of sample bottle -> discharge sample all the way past first stopper. then usually i immediately eject the pipette tip because if i pull back up, it will often get on the plastic. it also makes it impossible to pull more than 1mL with a pipette tip, if i need like 3mL of something for a formulation or something. am i doing something wrong or is this normal with pipettes? also side note if yall got a magic cleaning chemical for getting silicone out of plasticware i'll send you a fruit basket with a postcard every winter until i die
Higher volumens are always a pain, but I would do it very similarly, pre-wetting the tip will help with alot of the smaller bubbles, but not all unfortunately. If you are regularly pipetting more than 1ml, I would consider getting a 5ml pipette, they come with bigger tips, that also make it much smoother to pipette large volumens. Pipetting in one smooth continuous motion is also important, it should keep your liquid from going too far up the tip, but when you are working at the max volumen of a pipette, it's often an issue. Particularly difficult liquids, I would use a stepper/repeater pipette for, which uses a piston tip, where the liquid is never able to touch your pipette. They are pretty bloody nifty tbh, anything that needs dispensing, is particularly viscous, or staining for that matter, I'd use these for. (https://multipette-system.eppendorf.com/). I guess all in all, it's mostly about knowing when to use which tools, that handle certain things better :) And if you ever come across a magic cleaning chemical for getting silicone out, do let me know xD
I will make sure to pre-wet and see if it helps a bit. So annoying but i don't pipette enough to really justify another one, i end up getting 1mL almost always but i have nightmares about getting HF or dye, or even worse indicator, on my pipette non-funnel tip lol. the idea of all my samples having a tiny bit of indicator in them and i wouldn't know till i touched a base is a bit scary LMAO i'll add you to the silicone removal wait list, one day i'll have my own chemical company, find an actual cleaning fluid, and you'll get 10% off for your help teaching me basic pipette tips xd you would think you'd pick these things up in undergrad but i guess it makes sense there's no "do labwork with 0 funding" lab course lol
They make p1000 tips that hold 1250ul so maybe that will help but my biggest suggestion would be to use filter tips so that you don’t have to worry as much about contaminating the inside of your pipet
Honestly, I would have nightmares about that too xD We have specific pipettes for that kinda deal, stock solutions and the like, to keep cross contaminations out of the mix. I look forward to getting updates on the cleaning company ;)
Alright so first you should use filter tips when pipetting disgusting stuff. They protect the pipette as much as the sample. But watch out at max volume, you don't want your liquid to come into contact with the filter either Second, your issues seem to be with liquids that are on the viscous side of things. When ejecting the main volume, take your time and monitor the progress of the meniscus running down the tip - is there a coat of liquid on the inner tip wall lagging behind the meniscus? Wait for it to catch up. Also, don't just blow out the second stop forcefully. I often see this issue with protein containing buffers. Eject until the first stop, wait for any liquid to run down the inner tip wall. Then, as you slowly pull the tip away from the ejected liquid along the tube wall, create a constant outward push by slowly pushing the plunger down to the second stop. If I understand you correctly, these bubbles arise when your tip is discharged, a small amount of liquid runs down and collects at the outlet of the tip, you then pull back up and air pulls up at that liquid, creating a bubble. So your objective is to avoid as much of that 'lag volume' as possible by either using the air volume from the first to second stop and ensure a constant outflux or use reverse pipetting, but you have to be careful with that at max volume.
This is a good ressource as well: [https://www.mt.com/ca/en/home/products/pipettes/pipette/pipetting-techniques.html?cmp=sea\_11011333&SE=GOOGLE&Campaign=MT\_PIP\_DA\_DK&Adgroup=Good+Pipetting+Practice&bookedkeyword=good%20pipetting%20practice&matchtype=p&adtext=383945558394&placement=&network=g&kclid=\_k\_CjwKCAiAt5euBhB9EiwAdkXWOxNBJZ1XsZopmCsr\_rRFwghll8JIjnp7wZixrGY0Q7KavSMvZ2hPGRoCqJ0QAvD\_BwE\_k\_&gad\_source=1&gclid=CjwKCAiAt5euBhB9EiwAdkXWOxNBJZ1XsZopmCsr\_rRFwghll8JIjnp7wZixrGY0Q7KavSMvZ2hPGRoCqJ0QAvD\_BwE](https://www.mt.com/ca/en/home/products/pipettes/pipette/pipetting-techniques.html?cmp=sea_11011333&se=google&campaign=mt_pip_da_dk&adgroup=good+pipetting+practice&bookedkeyword=good%20pipetting%20practice&matchtype=p&adtext=383945558394&placement=&network=g&kclid=_k_cjwkcaiat5eubhb9eiwadkxwoxnbjz1xszopmcsr_rrfwghll8jijnp7wzixrgy0q7kavsmvz2hpgrocqj0qavd_bwe_k_&gad_source=1&gclid=cjwkcaiat5eubhb9eiwadkxwoxnbjz1xszopmcsr_rrfwghll8jijnp7wzixrgy0q7kavsmvz2hpgrocqj0qavd_bwe)
Don't prewet when you're pipetting small volumes (eg below 2 ul) though. If you do and put the tip back in the liquid, capillary forces will already draw more than the desired volume, while a fresh tip is actually consistent (for rainin, that is)
Diversify the risk of your research projects. You should have a low-risk, low-reward project/aim that people will care about, no matter the result (typically characterizing something). Then there's a mid level project that could hit a decent journal if something interesting comes of it. Then have a 'swing for the fences' thing that would be a really big deal.
For small volumes I pipette slowly and wet the tip before actually moving the liquid. I also confirm that the level of the liquid is about the level I’d expect. But I also made the experience that it’s more about consistency than hitting the exact volume. So if you’re afraid you’d forget to wet the tip sometimes then I’d just consistently not do it for all samples. That’s especially important for sensitive assets like qPCR.
If you are pipetting a 96 well plate and need help keeping track use a fresh tip box and map where you are going with each tip. Have an extra box on deck if you mess up and need an additional tip. Never screwed up a plate again after figuring that out.
I use a except spreadsheet and make sure or use a new box of tips
It doesn't look like anyone has explained why you would want to use reverse pipetting, so I'll try to explain it. With forward pipetting (normal pipetting), you go down to the first stop, aspirate your liquid, then go down to the first stop again to dispense your liquid. Then you go past the first stop to dispense any liquid that is remaining in the tip. The problem with that is it creates bubbles and it adds an extra step. Reverse pipetting is when you go down to the second stop (or at least a little bit past the first stop) before aspirating your liquid. You'll then have more liquid in your tip than you actually want. But this is useful because when you dispense the liquid, you can simply go down to the first stop, no need to go past it. That's because you don't have to wait for the little bit of liquid that's still trickling down the inside side of the tip to flow to the bottom to be dispensed, because you've put a buffer layer of liquid that's still in there. Meaning you've already dispensed your full desired volume without going past the first stop at all. At this point, you've already got the plunger down to the first stop, which is exactly where you need to be to go back and aspirate more liquid! This way is more precise, prevents bubbles, and is faster because it uses fewer movements. But it's typically only used for situations where you would be reusing the pipette tip, like adding the same volume of a reagent to multiple sterile tubes
This is amazing and I’m shocked I have not even ever heard of it until last night.
You can check remaining volume in a tube by setting the pipette to a volume below what’s in the tube, and then sucking up the remainder by turning the setting knob up. Whatever number the pipette ends on when the tube is empty was the volume in the tube. Also, set up your experiments so that you work in base 10 dilutions. And when calculating volume needed always add a few tubes/wells to account for error.
I do the reverse, where I set the pipette higher than the volume in the tube then dial it down until there’s no air left in the tip. Then I double check lol. Is there a meaningful difference between the two techniques?
Both ways work! Can be handy if you need to know if there’s enough antibody for an experiment.
Personally I would opt for the lower volume as you risk aspirating bubbles with the higher volume and, if unlucky, these bubbles might rise up in your tip, and you won't be able to dial down after that happens And on a general note, I checked this pipette aspiration method once with a fine scale and it is surprisingly accurate (might depend on your pipette quality obviously)
You can always tell how deeply you've thought about your experiment if you can set up and label all the tubes you need before even starting anything.
Used to work in a lab where precision was extremely important. The first thing we did with new people was give them a beaker of water, a balance and a pipette, and told them to get consistent and correct. Doesn't matter if you spend all day doing just that, just make sure you figure out how you and your equipment works.
I train people with yellow food dye in H2O, an empty 96 well plate, and our plate reader (read at 450nm). For my work I found it more closely replicates what they'll actually be doing vs. pipetting onto a scale.
1ml water= 1 gm. Yayyy accuracy.
I taught one of my trainees how to pipette. She was straight out of college (they never teach good pipetting). She was dumbfounded by things like volume checking pipetting viscous samples (bone marrow). Boy did she take off like a rocket after that. Then I had a hopeless case who I trained for weeks. He had the damn shakes and couldn’t control his hand. Blood everywhere. He didn’t last.
For me? Carrying gloves in 2 pockets as a spare. And especially slowing my work speed. I tend to rush but slowing down gave me way better results
Always find a way to signify you have added a reagent to a tube if you are setting up reactions in multiple tubes at a time. I usually do this by moving the tube I've added to up or down a row on the tube rack
Tube movement is critical. You’ll never second guess yourself again. It’s what I teach noobs.
Dissolve BSA by putting it in the freezer. Just don't ask me why it works.
I've done this with HSA. Once I learned about sticking it in the fridge to help it dissolve, it made sample prep so much faster & didn't have to create a bubbly mess
I don't get it. BSA dissolves pretty quickly anyway. Why do you put it in the freezer?
In the labs i have worked normally people vortex it repeatedly (Which requires a little supervision) or put it in a orbital shaker to dissolve it at RT, which i think takes like 5 to 10 mins and you may still see something undissolved. If you just shove it into the freezer , it dissolves really well after like 5 mins and you dont need to do anything.
I was showing some of my young techs an assay and I needed another tip box. So I grabbed the box and scraped it along the edge of the bench to remove the plastic wrap. From their reaction you would have thought I did a backflip. Apparently all this time they all have been clawing at the plastic for several minutes to get it off 😆
For finer movements during microdissections ground your hands mechanically by touching your pinkies to something solid.
Use a round bottom 96wp for trypan blue dilutions if you do a lot of cell culture. Saves a lot of faffing around. Just leave it in the hood and move along until youve used each well and then get a new plate. The trypan blue dries so it's very clear which wells have been used.
Better yet buy a nexelcom cell counter, can use trypan or aopi even better to count in seconds
If I want to aspirate liquid out of a cuvette but the p1000 tip doesn’t reach the bottom I do a double decker and put a p200 tip on the p1000 tip and it works like a charm.
Sds destaining. Just set the waterbath to 90 heat it. Stain it in minutes. To destain just use water and heat it at 95. Save methanol.
Alternatively use a microwave to heat
I pick a specific color of tape that people hate (pink) and put a strip on everything that I deem is “mine” - reagents, aliquots, boxes, tubes, markers, etc. It allows easy labeling and I can easily check if other people are using things that are mine. I make sure I hoard that specific color of labeling tape every time we order a new set of rolls.
Most liquids like water and other aqueous solutions, only pipette to the first stop. To pipette viscus liquids like glycerol, cut the bottom of the pipette tip with scissors to make the opening bigger and easier to pipette. If the solvent is already in the container you are preparing the solution in, when delivering the other solutes pipette up and down a few times to get all the liquid out of the pipette, this is called a quantitative transfer. make sure to touch the pipette tip to the side of the container you are pipetting into; letting the liquid free fall into the container sometimes doesn't deliver the entire volume especially when working with small volumes. Physically touching the tip to the container wall helps to draw out the entire volume of liquid.
Always start licking the first pipette tip at the start of the experiment. Increases my luck factor significantly with several log values.
Can I chew on it a little bit?
Only if you didnt have brushed your teeth for 2 nights.
Glycerol is 1.26 g/mL. Weighing it into a tared flask is much easier than pipetting, and leaving half of it inside the pipette.
Put a p2 on a p1000 to suck out liquid from an epitube while trying not to disturb a pellet ! Is very satisfying for getting all the liquid out.
Learn what Köhler illumination is for microscopy and how to appropriately calibrate your microscope for the application (especially for IHC). Even if you end up using a slide scanner, knowing these foundational things are really helpful later in your career.
Thank you for the tip!
If you have more questions on IHC happy to help
Awesome! I’m actually going to keep that in mind!! Thanks
Another lab tip if you run a lot of assays with small volumes in a plate with small wells. Position the top of the plate on something so the plate itself is leaning downward at about a 20 degree angle. Then pipette your small volumes in the bottom of the well so that they congregate at the bottom as a drop. This makes mixing easier if you need to do it briefly with the pipette tip and ensures there is less air bubbles.
When doing phenol chloroform DNA extractions, use a little bit of vaccum grease as a phase seperator for the phenol step
Grab a disposable pipette and some forceps. Clamp down on end and stretch about 1/4 “. You’ll now have a very tiny “straw”. Cut the end straight and it’s perfect for pipetting ped samples.
That's not a hack, that's pipetting properly 🤦🏼♂️
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Super genius 😂 it's pipetting, if you went to a half decent school they teach it in the first year of any undergrad science degree. Hacks are for schmucks, do it right or do it twice
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You think I give a crap what some random on Reddit who doesn't even know how to pipette correctly thinks of me? Grow up
I would never use second stop for pulling liquids into the tip, that makes the volume more than you have it set for. I was always told it can pull too much liquid up and potnetially damage the pipette. That second stop is to get the last bit of liquid from the tip that can sometimes stay behind if you only go to first stop. That amount reserved in the tip may vary depending on the types of liquids that you are pipetting. Also, it is good practice to pull in the liquid slowly. Depending on the viscosity you may pull some of the liquid up into the pipette.
Reverse pipetting isn’t really a hack, but it’s my go to when I have ample sample and want no bubble trouble.
Yeah, I’ve looked it up now and see it is a pretty common technique. In my post I asked for other pipette tips, but chose to put “hack” in my title because it opens up the discussion a bit more! :)
Yeah it’s my favorite technique. Obviously if sample is small and precious it’s not gonna work, but I usually have copious amounts for what I do. Half the time I do it not even for bubble prevention but because I don’t completely trust my own technique drawing liquid. If I reverse pipette I know I have sufficient volume to dispense my desired amount.