Tuesday, November 23, 2010

My Research Explained: DNA Tethering

It's been a while since I've posted on my research. Things in the lab are going smoothly and I'm taking my time with the data. My project is complicated and there is a lot to take into consideration when dealing with DNA unzipping (which I could incorporate into a future post). We've built a nicely working optical tweezers and now we are gearing up to unzipping real DNA. In order to take data efficiently and reliably I need to make sure that I can consistently create DNA tethers and in this post I will talk about how I do this.
When tethering is working, meaning we can get DNA to stick to a surface (glass in our case) it looks like this:

DNA attaches to our glass surface because of the interaction of an antigen and an antibody. We attach the antigen (digoxygenin) to our DNA molecule and the antibody (anti-digoxygenin) sticks to glass nonspecifically. Since we cannot see DNA in our microscope, because it is too small, we use larger particles (polystyrene spheres) to visualize proper DNA tethering. The spheres stick to the DNA due to another interaction between two molecules; biotin which is on our DNA and streptavidin which is on the microsphere. Below is a close-up of the setup:
The setup sounds simple enough: anti-dig sticks to glass then DNA sticks to the anti-dig and then beads stick to the DNA. Piece of cake. Actually not really. While it sounds simple enough there is a lot to take into account when setting up the experiment and it turns out a lot of it may not be well understood, but is done because it works and everyone does it that way.

Before I start the experiment, I set up my sample by constructing a microfluidics chamber. It is literally two glass slides separated by two strips of double-sided tape. The tape is spaced about 5mm apart creating a chamber that can hold a volume of 6-12ul (depending on how far about the tape is spaced). 
Next I get my samples ready. I need: anti-dig diluted in PBS (a buffer), diluted DNA, a washing buffer (I use a made up one Koch calls popping buffer), and diluted microspheres.

  1. First I flow the anti-dig into my chamber which is easy. I want the anti-dig to get near the glass and stick to it, which it does naturally. 
  2. From here on out, I have to be careful when flowing solutions because I rely on capillary forces to push solutions into the chamber. If the flow is too fast (force is high) I can damage my sample and if the flow is too low then I'm working inefficiently.
  3. Next I flow some washing buffer. The idea behind this step is to remove any free floating anti-dig. Since the DNA binds to anti-dig, I want to remove as much of it as possible to prevent DNA from sticking to whatever is not bound to the glass.
  4. Next I flow in my DNA. It turns out that different concentrations of DNA affect the appearance of tethers. Not enough DNA will be revealed because there will not be very many microspheres near the surface. Too much DNA will make the microspheres look like they are stuck. The perfect amount will yield a good stuck bead to tethered bead ratio (more on this later).
  5. Next I flow in more washing buffer to remove any free DNA and free anti-dig.
  6. Then I flow in the microspheres. While they are diluted, they are still really concentrated compared to what I want in the final analysis.
  7. To finish it off I flow washing buffer one more time to remove any free particles and remove most of the beads.
Now it's time to look in the microscope to analyze the results. There are 3 kinds of bead motion to look for. There are stuck beads which have no motion, there are free beads which move very randomly in the sample and move in and out of focus, and there are tethered beads which move randomly but are confined to a limited area (limited by the length of the DNA it is attached to). When only one DNA is attached to only one bead you get perfect tethered particle motion. Unfortunately more than one DNA molecule can attach to a single bead and this reduces the motion of the bead. If there is too much DNA, there can be a lot of molecules attached to one bead and the result appears to be a stuck bead. This next video shows differences in tethering efficiency.

Not only do you have to worry about the concentration, but there are a number of other factors involved. The effect of the washing buffer on the DNA tethering is not well understood and would take a lot of time to study what happens. Since I know tethering works with what I have, I'm just sticking with it. Another factor is the beads themselves. We use 0.5um beads. These beads are so small that they tend to clump up. I need to separate them somehow and my best tool is to sonicate them (a machine that uses ultra high frequency sound waves to clean various things). It isn't perfect but most of the time it works well enough.

So right now I'm at the stage where I'm trying to figure out exactly what will make DNA tethers the best so that I can repeat that recipe hundreds of times to produce award winning experiments. I'm taking it slow so I don't miss anything. I'm really close folks and hopefully next time I come to you about my research it will be about something truly exciting and ground breaking!

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