I was wondering if someone could shed some light on how to prove that a
protein or a complex is binding single-stranded (ss) or double-stranded (ds)
DNA and not just sticking to it when in pure form under given conditions. I
did gel shifts with a protein I am studying right now and ssDNA under
published conditions (Tris pH 8, Mg, ATP, ATP regeneration system).
According to published data that are quite scarce, the protein should
hydrolyse ATP in ssDNA-dependent fashion. Everybody seem to believe that
ssDNA binding reaction thus required ATP hydrolysis. However, I found out
that I can see the shift of almost the same intensity without addition of
ATP and Mg (even in presence of 2mM EDTA and no Mg). Moreover, I am totally
flabbergasted that DNA binding occurs even at 0-4C. Am I dealing with DNA
sticking rather than DNA-binding? Is there a way to find out if DNA binding
is real? The protein is not binding DNA in the sequence specific manner, so
adding excess "other" DNA or poly dIdC would not compete off non-specific
Any suggestions highly appreciated.
In article <FIa0b.38$[Only registered users see links. ]>, "EK" <[Only registered users see links. ]> wrote:
I don't know, Emir, to me sticking and binding is all the same thing. If
low affinity "sticking" has physiological significance, it's a perfectly good
binding in my book. Just measure Kd with all the approriate controls and
see if it all makes sense. DNA is easily to immobilize on an insoluble support,
so measuring your protein in the sup/pellet will do.
I am confused... If there is no sequence specificity and the binding is truly
to DNA then _any_ DNA should complete - isn't it?
something analogous to rnase protection assay comes into my mind: If your
protein binds ss DNA in a specific non-sequence specific way :-)), and you
digest away the non-bound part of the DNA, you should get some sort of
unique length DNA, you might be able to demonstrate these uniform size
fragments, when you label your DNA in a certain way (radioactivity might be
simplest). Maybe also non radiactive but sensitive detection like capillary
electrophoresis does the job.
In case of dsDNA, cloning the recovered blunt ended / blunted DNA and
determining the size and sequence of the inserts might give you lots of
If you have an idea of the size of the these fragments, by adding
synthetical oligos of different sizes each to your protein and measuring
the recovry, you could be able to determine the size dependency of the
effect and define a minimum length of ssDNA that will be bound.
Incubating the protein with a mix of oligos and doing MALDI/TOF or similar
procedures might be another possibility.
Fluorescent oligos and a suitable DNA sequencer also could do it.
"Tom Anderson" <[Only registered users see links. ].ac.uk> wrote in message
news:[Only registered users see links. ].ac.uk...
I might try heparin to copete off DNA. However, we use heparin
chromatography to purify the protein, so heparin will definitely bind the
protein. Most probably heparin will effectively compete off DNA.
I know from my recent experiments that salt at as low as 50mM concentration
will inhibit binding, meaning that possibly the process has some hydrophobic
component. Would that mean sticking vs. binding, I don't know yet.
Interestingly, iron (II) inhibits binding completely at 1 mM. Strange
behavior with Zn: 0 - 0.5mM gives too bands, and the lower MW band
disappears at 1mM and above. Ni - slight inhibition. Supposedly, reaction
should depend on Mg or Mn, but I don't see much difference between +Me
and -Me (even if +EDTA 5mM). Supposedly the reaction is ATP dependent, but I
see the shift at 0mM ATP. And, again, I see binding at 0C, too, although it
is a bit weaker. That "weaker" does not allow me to call the whole binding
a"sticking", although no dependence on Me and nucleotide cofactors speaks in
favor of bogus binding.
I just thought, when you have non-specific interaction, it either might be
the phosphate/deoxyribose backbone or some base feature in general (less
likely, probably). And if it's not just "DNA curling around your protein",
there should be a more or less defined length of DNA attached to your
protein, which also makes the DNA less accessible to appropriate DNAse.
Then you should see a more or less sharp peak when you analyze the size of
this nibbled DNA. I'd expect an unsharp peak since electrophoresis
resolution of DNA depends on size in basepairs, molecular weight and base
Yes and no. If you analyze enough clones and you really don't have sequence
specific binding, you should see a uniform distribution of base
compositions (i.e. having the same base distribution as the sequence you
start with in your probe - you might align your sequence data to your probe
and look at the distribution!!). I thought, sequence unspecifity is a part
of your hypothesis and this experiment might prove it.
Can you purify your protein on heparin agarose? When heparin competes with
DNA, then the binding is very much likely due to charged interactions, i.e.
the sulfate / phosphate groups. Then also phosphocellulose and
phosphotyrosine agarose chromatography should give similar results.
If the interaction is hydrophobic, you might be able to elute the protein
from a HIC column with short chain length polystyrene or similar (in
general a short polymer with aromatic side chains but uncharged).
It's just a shot in the dark, I'm afraid, but have you tried to see what
would happen if you biotinylated your DNA, bound it to streptavidin-coated
magnetic beads, and did a binding and elution series under various
conditions? It might make no difference at all, of course.