How much DNA is on the beads?

The NanoDrop tech support person said that the styrene beads wouldn't hurt the NanoDrop spec, and agreed that light scattering might be a problem.  It was, and that combined with the detection threshold of the Nanodrop meant that my measurements didn't give any evidence of DNA on my beads.  So today I used the PicoGreen assay to look for DNA on the beads.  It's much more sensitive, and not bothered much by light scattering due to the beads.

But first I should describe what my samples were and how I made them.  I incubated some 1.26 µ streptavidin-coated beads with a diluted solution of my biotin-labeled DNA, diluted because I didn't want different beads binding to the two ends of a fragment, and I didn't want steric interference by the DNA on the beads.  I incubated the beads with the DNA for 30 minutes, gently mixing at 37C on our roller wheel.  Then I pelleted the beads, and washed them twice with 1.0 ml of TE, each time rolling the beads plus TE for 10 minutes, and resuspended the washed beads in 100 µl TE (call these Beads1).  I also added another aliquot of beads to the DNA solution I'd already used with the first beads, and put these beads through the same incubation, washing and resuspension steps (call these Beads2).

Beads1 and Beads2 had very similar DNA concentrations, about 250 ng/ml.  This isn't very much DNA (but see below), but because they're the same I know that the low binding isn't because my biotin-labeling failed.  If the labeling had been the problem, then Beads1 would have had little DNA because they had bound up all the labeled DNA in the tube, and Beads2 would have had much less DNA. (I could check this by incubating more beads with the same DNA sample.)  Instead the low labeling may be because of the amount of strepavidin on the beads, or its reduced accessibility once bound DNA fragments are getting in the way of other DNA fragments.

So how much DNA is this per bead?  Here's a very back-of-the-envelope calculation:  The bead concentration in the resuspended Beads1 and Beads2 preps is about 0.1%, assuming that no beads were lost in the washing steps.  Let's consider 1 ml of Beads1 (or Beads2), just because it makes the arithmetic clearer.  With 0.1% beads, 1 ml of Beads1 solution is about 1 µl of packed beads (and yes, that's about how big the pellets appeared).  The beads are about 1.25 µ in diameter, and 1 µl is a cube that's 1000 µ on each side, so 1 µl of packed beads is a cube with about 800 beads per side, or about 5x10^8 beads.  At 250 ng/ml, the same ml of Beads1 contains about 250x10^9 kb of DNA (using Rosie's universal constant of 10^18 kb/gram of DNA).  The average fragment size of EcoRI-cut H. influenzae DNA is about 6 kb, so this is about 42x10^9 fragments.  I conclude that the average bead has about 85 DNA fragments bound to it.  That's pretty reasonable for my experiments, so I can go ahead and use these beads and this DNA to test cells binding to DNA on beads.

I also measured the DNA concentrations in the two washes from each aliquot of beads.  The first washes had about 20 ng/ml DNA, and the second washes had fluorescences not significantly higher than background, so I know that the signals from Beads1 and Beads2 were due to bound DNA.

One control I didn't do was to make a standard curve using known amounts of DNA mixed with 0.1% beads.  I should try this tomorrow.  I've also saved the samples I measured, and I'll also try reading them again tomorrow using the high-sensitivity setting of the plate scanner.  (Later - I was wrong; there is no high-sensitivity setting.) That's if I can figure out how to do this; the scanner software is very non-intuitive, and so far I've spent most of my time trying to find files I thought I'd saved.