Whatever molecular biology techniques you use, at some point you will have to clean up your DNA samples to remove things like buffers, contaminants and nucleotides from you precious sample, so that you have perfectly pure DNA for your downstream experiments.
Magnetic beads are one DNA cleanup option. They are simple and effective—and their reassuringly macroscopic nature is a welcome change for molecular biologists!
But even this seemingly simply technique can go wrong and deliver sub-optimal DNA end product. To help make sure that the simple stays effective, here are my seven tips for getting the best results from DNA cleanup with magnetic beads:
Don’t Freeze Your Magnetic Beads
Never freeze your beads. Unless specified, magnetized beads can never, ever, be frozen. Beads should always be kept at 2 to 8ºC. Freezing and thawing may cause cracks on the surfaces of your beads. And this can lead to sample contamination—the opposite of what you want.
Bring Your Beads to Room Temperature
While your magnetic beads should be stored and kept at 2 to 8ºC, you need to bring them to room temperature before you use them. So plan ahead and make sure to take your beads out of storage 30 minutes before you need them. This amount of time will give the beads time to warm up.
Homogenize Your Magnetic Beads
When you remove your beads from storage you will quickly realize that your bead slurry is not homogenous and the color is not even. Instead you will see a brown bead precipitation at the bottom of your tube. As stuff in molecular biology labs go, magnetic beads are huge! And, therefore, they unsurprisingly settle during storage. However, you need a homogenous bead slurry, so vortex your beads immediately before using them. Make sure that you vortex the beads until the slurry is an even color, and all your beads are in suspension. If your bead suspension isn’t homogenous, you risk endangering your protocol; either your current one or the following ones, if you keep using the same beads. And this is because…
You Must Know Your DNA-Bead Ratio
Protocols are specific for a reason and exact volumes matter. Bead input is directly correlated to the amount of DNA that you should obtain. Too many beads and you might get contaminants. Too few beads and you might lose product. So when you pipette your beads, make sure to clean up any droplet hanging on the outside of your pipette tip. And, of course, make sure to use good pipetting practices, including regularly calibrating your pipettes.
Pay Attention to Your Magnetic Stand
Make sure you choose a magnetic rack appropriate for what you are using—whether you are using 8-strip tubes, a 96-well plate, or a MIDI plate. The washing protocols usually include a lot of “remove supernatant and discard” steps. Losing some beads in these steps is normal, but you want to minimize bead loss—because, of course, your sample is attached to the beads! And, I am sure that you share my fear of losing sample during washes. So, use a well fitting magnetic rack.
Besides the size, racks vary in their magnet placement. Magnet placement affects where your beads will aggregate—whether it is at the bottom of the well, to the left, to the right, or in a ring conformation. If your protocol doesn’t specify which stand is best, you can choose based on your personal preferences. Just pay attention to what you are using! You will want to pipette away from the aggregated beads to minimize accidentally sucking your beads up. And try to find a stand with good magnetic strength.
Of course you know this, but here is my experienced pipetting advice: When you are aspirating your washes:
Keep your fingers steady. This is not a time to rush things or the day to have an extra shot of espresso in your latte.
Don’t aspirate too quickly. Go slow, as the strength of the aspiration can overcome the magnetic force and you can suck the beads on your pipette tip.
Monitor Your Loss
If you want to know just how much bead (and sample) you’re losing, then I advise you to keep a “waste well” for each sample. In other words, collect all your aspirated washes. At the end, you can put this waste well on a magnetic stand and actually see how many beads you are losing with each wash step. This can help you clarify why some samples might work better than other. Not qualitative but still helpful.
If you have good bead practice, a steady finger, and a focused eye, I guarantee you will keep bead waste to a minimum and increase retention of your purified sample. Happy Washings!
You might be proud of your pipetting skills (if not, check this article on how to stop pipetting errors from ruining your experiments) and be churning out data faster than a liquid handling robot, but beware… you might also be pipetting yourself out of a job. I almost did. Pain due to pipetting is common. […]
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