Koch Lab:Protocols/Microsphere-DNA tethering/Glass, dig, biotin, microsphere, 4kb DNA/Cleaning glass

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Even without cleaning the glass surfaces, we have been able to achieve decent tethering (high tether density relative to stuck beads) following the protocol on the parent page. However, a standard and repeatable cleaning method is bound to provide more consistent results and is likely desirable (although, the cleaning procedure may not necessarily increase the tether density, I can't promise that). There are several different methods we are familiar with, and others we are not. In order of familiarity below:

Crazy Windex Method

In graduate school, Steve mostly desired consistent results. He reasoned that Windex was good at making glass appear to be clean, and is cheap. He also reasoned that residue left behind by the Windex was likely more consistent than whatever gunk is on the glass to begin with. He was in a hurry and didn't like the sonication in ethanol / sonication in water method. So he did this with very consistent results:

For each glass surface (one coverglass, one slide):

  1. Spray with a squirt of Windex
  2. Scrub with a kim wipe
  3. Blow dry with compressed nitrogen or clean air
  4. Rinse with nanopure water (no scrubbing)
  5. Blow dry with air
    • Repeat above two steps (water rinse + blow dry) for a total of four rinses

The coverglasses appear clean to begin with usually. The slides typically have cloudy patches to begin with. Both appear clean after this procedure.

Sonication in Ethanol then DI Water

I saw people get consistently good results with this method, and I also got good results myself but abandoned it in favor of quicker Windex method above. So, I am not sure of the details of the protocol anymore, but it's something like:

  • Get a tray that allows you to stack individual coverglasses in a row so they are not touching each other.
  • Get a similar tray for slides
  1. Put tray in a beaker submerged in Ethanol
  2. Put beaker + submerged tray in bath sonicator and sonicate for a long time. Like 15 minutes I think.
  3. Remove tray from ethanol and rinse with lots of DI water.
  4. Put tray in a beaker submerged in DI water
  5. Put beaker + submerged tray in bath sonicator and sonicate for a long time, as above.
  6. Remove tray, and rinse with lots of DI water.
  7. Dry tray in an oven.
  8. Store in container protected from dust.

Sonication in Water and Alconox

It is good to keep a 1% solution of alconox around the lab for cleaning in general and this protocol makes use of that. I'm sure you could use some other good surfactant or cleaning product as a replacement. You will also need a tray (like above) to hold cover glass and slides.

  1. Fill sonicator about halfway with water and add a little Alconox to the water. Stir solution.
  2. Submerge tray with glass.
  3. Turn on sonicator and sonicate for at least 5 minutes.
  4. Remove tray and rinse slides with lots of DI water.
  5. Blow dry
  6. Store in a container protected from dust.

UV-Ozone Cleaner

At Sandia, I had access to a UV-Ozone cleaner. I didn't use the method enough to know whether it was consistent, but I think it is pretty promising.

  1. Start with a few coverglasses and slides (a few that you will use in the afternoon)
  2. Place them in the UV-Ozone cleaner
    • In the one I used, we put them on a tray, on top of slides that remain in the tray
  3. Clean for 15 minutes
    • You can test the effectiveness of cleaning & how much time is needed by observing how well a drop of water wets the surface after cleaning.
  4. Make not of the clean sides (facing up) when you remove, and make double-stick tape chambers right away

Other methods

  • Piranha wash. This one seemed too nasty for me to deal with. But a chemist could probably show you how to do it and assure you that it's no big deal. Some mixture of hydrogen peroxide + sulfuric acid.