Investigating DNA - Mercury Binding Interactions using a Fluorescent Dye

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Investigating DNA - Mercury Binding Interactions using a Fluorescent Dye


In my last post, I demonstrated that using a fluorescent dye that can insert between DNA bases can help me learn more about the intricacies of the DNA - mercury binding system including binding stoichiometry and affinity! Let's get right into this very colorful biochemistry!

What Dye did I use?

I mentioned it briefly in my last post, but for these experiments, I used an intercalating fluorescent dye called Thiazole Orange. Intercalating just means that the dye is able to insert itself in between the bases of a DNA molecule. The rationale behind using this method is that as our single-stranded DNA molecules in solution encounter mercury ions, they form a hairpin structure or some kind of secondary structure around the mercury ion per our hypothesis. Thus, the ssDNA's own bases then loop around and become in close contact with each other. Close enough for this dye to attach. 

But how does the insertion of the dye actually result in the fluorescence of light? Well, let's start with just our Thiazole Orange (TO) in solution, without any DNA nearby. When it is blasted with light at 510nm (its specific excitation wavelength) the electrons in the molecule are elevated from their natural ground state and into an excited state where they occupy higher energy levels. However, there are specific chemical bonds in TO that allow for a lot of rotational motion in the molecule. So, when it isn't attached to any DNA strands, the molecule literally gets rid of all the excess energy by simply rotating and rotating and rotating in solution. 

Now,  what happens when attaches to the bases of a DNA molecule? Well it turns out that the rotational bond is completely restricted by the DNA bases and there's now no way for the molecule to get rid of all the excess energy via rotational motion. But there's still a way. The molecule is now forced to release the energy via expelling photons, i.e., light. This light is released at the highest intensity around 535nm. This is called emission and a molecules emission wavelength is always a larger value than the excitation wavelength.

What did I use the Dye for?

I used the dye to perform spectrofluorometry experiments to ascertain two primary facts about the DNA - mercury system: Binding stoichiometry and binding affinity. I'll start with the stoichiometry first but before I do I want to briefly outline how I set up the samples in general. Each sample, in general, contained a calculated amount of 6uM DNA, a calculated amount of 1mM mercury (from my mercury perchlorate stock), an excess concentration of my TO dye, and a backfill of HEPES buffer as the diluent.

Determining Binding Stoichiometry

To determine this very important property of the DNA - mercury system, we needed to construct something called the Job plot (named after the guy who perfected the method). A job plot uses what's called the method of continuous variations, which is a fancy way of saying that you're constantly varying the concentrations of both the DNA and the ligand (in this case mercury) over the course of the experiment. 

A total concentration was determined to be 10uM so that means the total concentration of both DNA and mercury must be 10uM. So that means combinations of 1 and 9, 2 and 8, 3 and 7, and so on. This is recorded as the mole fraction (mol frac) of mercury, so 0uM mercury was 0 mol frac mercury and 10uM mercury was 1 mol frac mercury.

So each sample contains a varied relative amount/concentration of both the DNA and the mercury. At this point, each sample was then placed in a spectrofluorometer and the maximum fluorescence was measured and recorded. The maximum fluorescence arises at the wavelength at which TO releases light the most intensely. Again, as a reminder, this is because the TO is binding to the DNA molecules as they are interacting/wrapping around the mercury ions.

The next step is to plot maximum fluorescence on the y-axis and the mole fraction of mercury on the x-axis. And here's the interesting part. The place where the maximum fluorescence on the Job plot itself reaches a maximum is correlated to the binding stoichiometry of the system! Job plots usually show a clear ascent/descent with a clear peak indicating the maximum. For example, if the peak is at an x-value of 0.5 mol frac mercury that means the binding stoichiometry is roughly 1:1! And if the peak is at a value of 0.75 mol frac mercury like the graph below shows, that means the stoichiometry is 3:1 mercury:DNA! Based on our Job plot, the peak max fluorescence was at about 0.77 mol frac mercury which suggests that 3 mercury ions bind to each strand of DNA!

Determining Binding Affinity

After finding the binding stoichiometry to be approximately 3 mercury ions to 1 DNA molecule, it was time to move on to investigating binding affinity i.e. how tightly the mercury binds to the DNA. The way to do this is to make what's called a titration curve. 

A titration curve (in the context of biochemical binding studies) arises from an experiment where either the DNA or the mercury concentration is kept constant and then the other is added incrementally in very small amounts.

In this case, I kept the DNA concentration constant at 6uM and added mercury to the DNA solution. After each small addition of mercury (3 - 6 microliters at a time) I measured the maximum fluorescence like I did when assembling the Job plot. However, I plotted the maximum fluorescence on the y-axis with the concentration of mercury on the x-axis, instead of the mole fraction.

The shape of a titration curve is much different, as can be seen below. It has a sigmoidal shape to it and can therefore be fit by a sigmoidal curve, which is the red line passing through all the data points. But it's not just a curve; specifically the data points are fit to the Hill equation which solves for two variables: The dissociation constant and the Hill coefficient or cooperativity constant. 

The dissociation constant is defined as the concentration of mercury ions at half of the maximum fluorescence measured in the system. It's the value that describes how much affinity the mercury has in binding to the DNA, but it's an inverse relationship. The higher the value, the lower the binding affinity. Our value of 14.4uM signifies a moderate to borderline weak binding affinity which is actually in line with the literature in terms of metal cation binding to DNA.

Additionally, the cooperativity constant describes how cooperative the system is. If the value is >1 then mercury binding encourages even more binding after that point. If the value is <1 then mercury binding actually inhibits further binding of other mercury ions. If the value = 1 then there is no preference either way. The determined cooperativity constant of approximately 3 suggests that when the first ion binds to the DNA, it very much encourages other ions to bind to the DNA down the road. This is very useful information for our DNA - mercury system!

One Last Interesting Note

One interesting thing about our Job plot is that there was a kind of weird "lag phase" where the fluorescence didn't really change at all until after 0.5 mol frac mercury. We're not really sure why this is but it's important to note that the titration curve is actually in agreement with this as well. 

In the Job plot, binding doesn't seem to happen until a 1:1 mercury:DNA ratio. Then, in regards to the titration curve, remember that I said I kept the DNA constant at 6uM? Well the fluorescence doesn't seem to increase here much either until about 6-8uM mercury. Again, this is close to that 1:1 ratio. So, for some reason, no binding is happening until there are equal amounts of both mercury and DNA. We still don't really know why that's the case! That's sometimes what's fun about science!

Moving Forward

During this whole process, I was able to move forward with my sample prep for the scanning electron images. In my next post I'll describe that process in detail and post the amazing images we ended up getting! It's incredible!

Also, as a disclaimer, I was unable to actually post the graphs I've been talking about since they may be included in a publication soon. Posting them online may result in the work getting stolen, which is not something I want to happen.

Thank you for reading and I'll see you on the next post!



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