Author: Prof. Andresen

We picked up this summer almost exactly where I left off my training with Savannah. Since I first started, the organization of the lab had been bothering me. Especially the bookshelf, I just did found the entropy to be too high. This is no longer the case! A very thorough cleaning has left us with several empty drawers, an alphabetized bookcase and an alphabetized cabinet of  samples. Additionally, I prepared all of the samples required to calibrate the ICP and the calculations required to do so. This process included massing several solutions and then diluting them to a specific concentrations. After I had done this Professor Andresen believed that I was qualified enough to write my very first Wiki article on the subject. The process of creating my very first wiki page was exhilarating, it feels almost like a child to me now. Today, Wednesday, I was almost able to set up the ICP machine by myself. My goal for today is to run the ICP on the samples that I prepared yesterday. Below is a picture from my notebook of the theoretically calculated masses of the samples I prepared and below the theoretical masses are the actual ones.

This week was my final week of research this summer. I was very busy all week trying to get as much done as I could before I had to leave. On Monday I reran the DLS for the new cleaned DNANP samples and saw the same dramatic increase in size following cleaning. We still do not know why this phenomenon is occurring in the new samples but was not seen the original (10 ug/mL) sample. There are not any obvious signs of too much aggregation in the salt-TE cleaned samples so it is unclear why the phenomenon is occurring at all.

                                  

I didn’t have ICP gas to run my samples from last week spin cleaning so instead on Monday I ran UV vis on the supernatant samples to confirm that the free DNA concentration decreased with each spin. The free DNA in solution plummeted after the first spin so I am confident that the spinning protocol is enough to clean the free DNA out of the solution leaving only bound DNA- NP complexes.

                       

Today is my last day in lab for the summer. In the beginning of June, I began by ordering a stock of whole chicken blood, and proceeded to purify this stock throughout the summer. For the past few weeks in particular, I had been working on running gels to digest the sample with the correct amount of micrococcal nuclease. This week, I correctly digested both of my samples, so they are now ready to be run in a column to isolate the mononucleosomes. In the fall, I look forward to further purifying my samples until I just have mononucleosomes, and running electrostatic experiments on them!

For the past several weeks, I have been working on “digesting” my sample, and running sample os varying degrees of digestion with gel electrophoresis. The first step of the process is to do a “trial digest,” in which different concentrations of micrococcal nuclease are added to the nucleosomes. Micrococcal nuclease effectively “eats” the DNA, slowing down when it approaches the histone core. A higher concentration of micrococcal nuclease will “eat” more DNA, so the optimal amount that will digest only the linker DNA is sought.

Next, proteinase K is added to the nucleosomes, which digests the histone proteins. There is now free DNA in the sample, and its length is determined by the amount digested by the micrococcal nuclease. Gel electrophoresis can be done on the variously digested samples to qualitatively see how long the DNA is for each micrococcal nuclease concentration (proteinase K concentration stays the same). The goal of this “trial digest” procedure is to determine what concentration of micrococcal nuclease digests just the linker DNA, so that all we are left with in the sample is the histone core and DNA wrapped around it. It is known that there is approximately 146 bp of DNA around a histone core (without the linker DNA).

It took several attempts to get a successful gel though. Examples of successful and unsuccessful gels are below.

This gel is an example of a gel that was not successful. It is a good gel in regards to the quality of the 10 bp DNA ladders (in the first and last lanes), but the samples are all trapped in the wells, rather than moving down the gel like the DNA ladder did. Such behavior could be explained by improper digestion, perhaps because of the micrococcal nuclease itself, its digestive medium, or other factors.

This gel is an example of a successful gel! Although the DNA ladder is not as clear as the gel above, the samples did not stay in the wells, and instead, moved down the gel depending on their length. It is interesting, though, that a majority of the digested samples have a length of approximately 300 bp (double what it should be!).

Because the 40 units of micrococcal nuclease seemed like the optimal concentration to digest the DNA, the sample was digested with this concentration, and compared in another gel to the prior undigested sample (see image below).

This gel contains the DNA ladder (not very clear), the digested sample, and the undigested sample. It is good that there is a stark difference between the digested and undigested sample…but where exactly is the undigested sample? Is there DNA there, is it over digested, or does the concentration of DNA in the gel sample need to be increased?

The goal this week was to clean all of the unbound DNA out of a stable DNA-nanoparticle solution so that we can quantify the amount of DNA per nanoparticle. Due to the results we saw last week we used 10 ug/mL DNA in the same 0.3 nM CTAB gold nanoparticles. The protocol we came up with was to spin down the nanoparticles and then do 3 washes with TE+10mM NaCl followed by 2 washes with pure water. While doing this I encountered some aggregation and incomplete pelleting. The aggregation was reduced after each spin indicating that the spinning was clearing out any unstable particles from the solution. The incomplete pelleting was combated by spinning the collected supernatant 1-2 extra times and collecting the extra pellet and a hopefully clean supernatant. The result of this protocol modification was 3 times as many spins. Each spin took 40 minutes so the protocol took two very long days to complete. The result was two DNA-nanoparticle samples a sample washed by salt-TE buffer and water and a sample washed only with salt-TE. More aggregation was seen after adding cleaning with the water indicating that either the salt or the TE buffer is integral in maintaining the stability of the nanoparticle complexes.

Failed gel

I started this week trying to successfully clean the free lysine out of my solution of lysine capped citrate gold nanoparticles. My repeated efforts to spin down the nanoparticle complexes without aggregating them proved ultimately futile. But while I was having a battle of wills with my uncooperative nanoparticles I learned that Professor Thompson’s lab had finally gotten a new supply of CTAB and had finally landed on a recipe that yielded CTAB gold nanoparticles of the proper size, shape, and concentration for my experiment. So I abandoned the lysine-citrate gold nanoparticle system and went back to my original system. I started by titrating my sheared DNA into the nanoparticles. The resultant UV-vis spectra showed a great deal of aggregation which was surprising.

The next day I made full samples of the 0.1 ug/mL, 1 ug/mL, 10 ug/mL DNA-CTAB AuNPs and measured their UV-vis and DLS against the control. Interestingly, the sample with the smallest concentration of DNA instantly aggregated whereas the solutions with more DNA showed little to no aggregation. The previous titration showed aggregation in all the samples because the severe aggregation caused by the low concentration DNA in the beginning could not be recovered by the addition of more DNA.

In order: Control, 0.1 ug/mL, 1 ug/mL, 10 ug/mL

This phenomenon has been reported before in a paper on DNA electrostatic interaction with DMAP AuNPs and is attributed to lower concentrations not being able to fully cover NPs resulting in a drop in charge that facilitates aggregation (Biver et al.).

This week I couldn’t really do much lab work because I was waiting for the lysine I needed to cap my citrate gold nanoparticles with. For most of the week I read papers on the lysine capping process and on the theory of DNA-gold nanoparticle interactions. On Thursday, I finally got my lysine, I made stock solutions, and I combined them with the citrate gold nanoparticles to initiate the capping process. I wasn’t able to clean the nanoparticles, however, as the students in Thompson’s lab are finishing up their research and need the centrifuges for the next two days. I went ahead and characterized the unclean nanoparticles though with UV-vis, DLS, and Zeta. Next week I will clean and characterize the particles and attempt to wrap the sheared DNA around them.