Washing beads free of unattached DNA

So a while back I attached some more DNA to my polystyrene beads, but I didn't get to measuring how much DNA because I was having problems washing the beads+DNA free of the unattached DNA.  I wanted to just pellet the beads+DNA in a microcentrifuge, remove all the supernatant liquid, and resuspend the beads+DNA in fresh TE.  Because the volume of the pellet should be less than 15 µl, repeating this several times should be enough to bring the concentration of free DNA below its initial value of about 50 µg/ml to less than 1 ng/ml, well below the expected concentration of the DNA attached to the beads and thus low enough not to interfere with measuring how much DNA is attached to each bead.

BUT, the beads were difficult to pellet.  Some of them formed a pellet at the bottom of the tube, but the rest seemed to just stick to the sides of the plastic tube, even though they were subject to a centrifugal force of more than 10,000 times gravity.  Because the spinning tubes are oriented at an angle to the centrifugal force, I tried my usual trick of interrupting the spin to rotate the tubes 180°C, but the beads still didn't dislodge.

With this strong force, I had expected that the beads would be packed tightly ino the pellet, but they weren't - they dislodged all too easily while I was trying to remove the liquid above them*.  And the beads that, during centrifugation, behaved as if they were stuck firmly to the sides also dislodged easily when I was removing liquid.  The overall result was that either a lot of the beads had to be removed with the liquid, or a lot of the liquid had to be left with the beads.

One way to prevent things from sticking to surfaces is to coal the surface with a silane.  But pre-treating my microscentrifuge tubes with a silinazing solution ("Sigmacote") didn't make a big difference.

Then I thought of a totally different solution.  The beads are about the same size as bacterial cells, which we routinely collect and wash by filtration rather than by centrifugation.  Maybe I could wash the unwanted DNA from the beads+DNA by filtration.

The photo shows what we use to filter our cells: "Analytical Test Filter Funnels" from Nalgene.  They're sterile, fit on a vacuum flask, and have cellulose nitrate membranes with 0.2 µ pores.  The liquid flows through, and the retained cells are washed by flowing through a washing solution.  The filter funnels snap apart to allow the filter plus the retained cells to be easily retrieved.  We recover the washed cells by simply shaking the filter in a flask or tube with some medium.  The funnel can hold 100 ml of liquid, but we never use the full capacity because the filter becomes clogged if we try to use more than 10 ml of cell culture at an OD of 0.25.

So I tested using a filter funnel to wash some beads.  The first test wasn't very successful; although the filtering and washing seemed to work very well, about half of the beads stayed on the filter even after washing with 1% SDS.  But I tried again, using 2.1 µ beads rather than 1.2 µ, keeping the vacuum weak, making sure that the filter was never sucked dry, and recovering the beads by gently shaking the filter+beads for 15 minutes.  This gave full recovery of the beads, in 3 ml of liquid. 

But I need to concentrate my beads as well as wash them, and 3 ml is much too dilute.  So I still need to test whether I can use filtration to remove most of the unwanted liquid.  Perhaps I can just pipette the 3 ml onto a small disk of membrane filter, with some absorbent material beneath.  Or maybe I can put the filter disk on the filter of another funnel.

*  In hindsight this makes sense; the beads are rigid spheres, not squishy cells, so they can't really be 'packed down'.  However my biophysicist advisor told me she had been instructed to not spin the beads harder than 3,000 times gravity as they would deform.  So I checked the ones I'd spun hard - they all looked spherical under the microscope, but the laser tweezers may be much more sensitive to deformation than my eyes are.  Filtering will eliminate this concern too.