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Dr./Mrs. Jekyll/Hyde has been teaching a new laboratory tech the tricks of the trade (or, as it turns out, some basic algebra), and in one of the posts mentioned something that we often don’t think about in science education but is critically important when you’re actually in a lab: having good hands.
This is important for a variety of reasons: pipetting solutions, filling the wells of a gel before running electrophoresis experiments, or even simpler things like not stabbing yourself with a needle or not dropping expensive samples. The shakes can be the difference between having to do experiments only once and wasting an entire week repeating them, with questionable results each time.
Not drinking coffee is, for me at least, not an option. Thankfully, my Ph.D. work involves very little that explicitly requires extremely good hands (mind you, pipetting solutions in a dark room under dim red lights* is annoying no matter how good your hands are…). I was not so lucky for my Master’s work, where, you may recall, I dealt with micropipettes.
Making the micropipettes was annoying for consistency reasons, but actually getting a micropipette into a flow chamber was the real fun part. Our flow chambers looked like this:
Brown tubing, having an outer diameter of ~160 microns, and an inner diameter about half that, was squished between melted Nescofilm between two No.1 coverslips, and was the means of inserting micropipettes into the active flow area.
The micropipettes, shown on the right, were tapered to ~1 micron at the very tip (the polystyrene bead held by suction at the tip is 2 microns in diameter), and for practical reasons were usually 3-5 cm long. They were connected to a length of thin tubing so as to allow for easy connection to a syringe, and the whole contraption was mounted on an aluminum clamp, with a separate clamp specifically to hold the micropipette in place.
As I mentioned above, I drink a lot of coffee, and while I don’t have a serious case of the shakes, I don’t have a particularly steady hand. Despite this, I would insert the micropipettes into the brown tubing by hand. We had a 20x optical microscope with two-dimensional motion on the microscope stage. I would mount the flow chamber to the stage, centre everything such that it was all in focus, bring a micropipette tip into the field of view by hand, and, once also focussed, try to move the stage so that the pipette tip ended up in the brown tubing.
This would (obviously) often result in disaster, as any time the tip of the pipette would touch the brown tubing it would be ruined, no matter how incidental the contact. For a while I was keeping a log of successful tips to ruined ones in my lab book, and the ratio got so depressing I had to stop. By the end of my Master’s though, despite my regular multiple cups per day, I had developed a couple of tricks to raise
the success rate. The most useful of which was to introduce water into the system.
Flushing the flow chamber with water would produce a large (under the microscope, anyway) bead at the opening aperture of the brown tubing, which, after Kimwiping it away, would leave a slight meniscus of water right at the entrance. When the tip of a micropipette was nearing the entrance to the brown tubing, the pressure difference in the pipette tip would result in rapid suction of water from the meniscus into the pipette. With much practice, rapidly moving the stage once observing water in the pipette led to, near the end anyway, maybe as high as a 50% success rate in getting tips into chambers.
While my Ph.D. work has it’s own series of issues (this week’s: overlap two 200 femtosecond pulses focused into a ~100 micron diameter area. Spatially is simple, temporally, less so), I am very happy I don’t have to do anything like that anymore. I think I’ll have another espresso to celebrate!
* I’m working on the most-obviously-photoactive protein you can imagine. Take a guess which!
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Biocurious is written by Andre Brown and Philip Johnson, since 2005. Content of the weblog is licensed under a Creative Commons Attribution-Share Alike 3.0 License.
Why do that when you can use a micromanipulator?
Mostly it was because of time, and the awkwardness of the mounts we were using. It was significantly faster (for me, others in the lab had their own favourite tricks) to do this by hand, and given that consistency problems existed with pipette tips to begin with, it felt to me a better idea to burn through a dozen in the span of a few minutes to get a good tip than to try each individual tip much more slowly.
It was also my opinion that the size of the tips made it very difficult to tell when they were in focus anyway, so even going the micrometer adjustment route, everyone’s success rate was poor.
That being said, you are probably right that there was a better way.
I do a lot of micropipetting, and I notice that my hands are much shakier anytime I’m hungry. For me at least, my steadiness has a lot more to do with my blood sugar level than with caffeine intake.
Relaxing my arms and hands is also important—if I’m nervous about the result of an experiment, I tend to grip the micropipetter with a ferocious intensity that makes my hands shaky. A few years ago I got tendonitis from too much pipetting, so I started making an active effort to hold the pipetter as gently as possible, and a pleasant side effect was steadier hands.
Why do U use the brown tubing ??
I was using it before and now I just put directly in-between the two nescofilms.
Not only is much easier but it makes also the system much stable.
Wilfried,
We used brown tubing so that you could re-use the same flow cells over again, even after a pipette tip might break or get clogged.
What kind of stability are you referring to? I can’t say we ever had problems with the flow cells because of the brown tubing being present.
Hi,
I had similar problems with flow chambers. You can check photos of our setup here:
http://users.ox.ac.uk/~jesu1458/malinauskas_bsc_thesis.pdf
Hope that helps,
T.
Philip,
The pipette is not as stable in the brown tubing as it is when it is firmly hold directly in between the Nescofilm.
W.
While that’s probably true, when using a micropipette to suction a bead on the tip to use in single-molecule force-extension experiments, the relevant parameter is how stiff the pipette is relative to the optical trap. I don’t recall the number offhand, but I do remember that the pipette was three or four orders of magnitude more “stiff” (in terms of an effective spring constant) than a bead held in an optical trap.
You also don’t have to remake a bunch of chambers any time your pipette tip gets clogged!