Friday, July 29, 2016

The End of the Line

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.

Then I ran ICP-OES on 10x and 1x dilutions of the supernatant and DNANP samples. A preliminary look at the ICP data revealed that the phosphorous concentration (2 phosphorus=1bp of DNA) was far below the detection limit even in the 1x sample. I then concentrated the remaining 10 ug/mL DNANP sample by 8x in order to hopefully get at least one measurement with detectable phosphorous. Even with the increased concentration the phosphorous was still undetectable. I will likely need to start from the beginning with a lot more sample so that I can concentrate the final sample by a larger factor but I will not be able to try that until the fall. With the remaining time this week I made and ran a new ICP calibration that covered sodium concentrations less than .500 ppm. This was applied to my water washed samples to hopefully determine whether any significant amount of sodium remained bound to the DNANP complex. I also reran the 10x and 1x dilutions to get more data. Today I have been working on compiling and analysing the data from the ICP. There seems to be a great deal of fluctuation and error in the ICP measurements which is concerning.


Friday, July 22, 2016

More Spinning and a Wee Bit O' Fun

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.

This was Sarah's last week so on Wednesday we took time out of the day to get lunch and ice cream. Since this took most of my day on Wednesday I did not try to start another spinning procedure until Thursday. I prepared 4 new DNA-NP solutions with new concentrations of DNA (20 ug/mL, 30 ug/mL, 40 ug/mL, and 50 ug/mL), characterised them and repeated the cleaning protocol with 3 washes of TE-10 mM NaCl and 2 of pure water. Similar to last week I saw much more aggregation following the water washes than with the salt TE. The UV-vis data showed very little shift in the spectra but a decrease in concentration and an increase in aggregation following the washes. The most interesting part of the characterisation data was that the six of the particles jumped from ~60 nm prewash to ~80-100 nm after being washed in the salt and the water. The jump in size was most pronounced in the lower concentration. The 50 ug/mL sample was back down at ~66 nm. This is interesting because no jump was seen in the samples from last week which could be due to subtle differences in the procedure or the fact that a different sample of sheared DNA was used. Next week I will run ICP which should give me more insight into the complexes.


Wednesday, July 20, 2016

July 20th - My Last Day in Lab!

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!

Monday, July 18, 2016

July 18th - Gels, Gels, and More Gels

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?

Friday, July 15, 2016

Endless Spinning

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.
I was going to start measuring the samples in the ICP but we were out of the argon and nitrogen gas. Instead I made 7 more sheared DNA samples, ran UV-vis on them and attempted to run them in an agarose gel. The UV-vis showed normal DNA spectra with DNA concentrations between 0.3-0.6 mg/mL. The agarose gel did not work and will need to be done again next week.
Failed gel

Friday, July 8, 2016

The Return of the CTAB

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.).

Biver, T. et al. “Analysis of 4-Dimethylaminopyridine (DMAP)-Gold Nanoparticles Behaviour in Solution and of Their Interaction with Calf Thymus DNA and Living Cells.” Journal of Nanoparticle Research 14.2 (2012): 1–12. Web.

Thursday, June 30, 2016

A Whole Lot of Waiting

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.

June 30th - Gel Electrophoresis Struggles

For the past week and a half, I have been doing gel electrophoresis to determine ideal concentrations of various chemicals to digest my DNA and histone proteins. Gel electrophoresis is a biological technique used to determine the size of the molecules in samples. For DNA, the technique is especially useful at determining how many base pairs a strand of DNA has. It is a mostly qualitative measurement, where samples can be compared to a known DNA ladder standard. Gel electrophoresis works by inserting samples into lanes of a gel, and applying a current through the gel and a buffer medium. Because DNA is negatively charged, it will travel towards the anode. Shorter DNA will travel faster though, because it is more easily able to maneuver through the maze-like holes in the gel.
Below is an image of the first gel I ran. The DNA ladders on the two ends are very crisp and visible – but there is a problem with the other samples; none of them moved. Such non-movement was likely because the digestions were not working. We have attempted different concentrations of our digestion chemicals, but have yet to find a solution. While we wait for more supplies to be ordered, I will begin making another sample of nucleosomes again from the 50 mL of whole chicken blood. Hopefully the gel electrophoresis problems will soon be resolved, and we won’t have this issue again in the future!

Friday, June 24, 2016

Black Flecks of Aggregation

I started out this week by trying to purify the DNA-NP conjugates that I made last Friday. Unfortunately, all of the samples except the 1 microgram/mL DNA sample showed significant aggregation after sitting at room temperature for the weekend. I chose to only attempt purification of that one sample but after just 2 spins in the centrifuge the sample had completely aggregated out. I tried the same concentration with slower spin speeds and longer times but the sample continued to aggregate out. Thinking that perhaps the conjugates were forming histone-like aggregates, I next tried digesting the "nucleosomes" with the micrococcal nuclease, an enzyme Sarah is using to isolate mononucleosomes, and then purify them. The result was a low concentration solution with lots of black flecks of aggregation that floated in solution and resisted pelleting following extensive centrifugation.
If you look closely you can see the black flecks of aggregation
 Unfortunately in the Thompson lab, they ran out of nanoparticles and the CTAB needed to make new nanoparticles so instead I turned my focus to shearing down the calf thymus DNA using sonication so it would be less likely to incite aggregation. I had attempted this in the spring but had not had any success mostly due to the fact that I had to use a water sonicator that is not optimised for DNA shearing. The protocol called for a probe sonicator. There is only probe sonicator at Gettysburg in the Biochemistry lab and I had been told it was broken. However, with nothing else to do, we decided to take the probe sonicator and try to fix it. Turns out the sonicator was not broken, it worked just fine once we plugged it in. So I ran the shearing protocol I found in a paper I read. The sonicator is quite loud. I only ran it at level 3, but higher levels probably require ear protection.

After I ran the shearing protocol, I characterised the sheared and unsheared DNA using DLS, UV-vis, and gel electrophoresis. The gel electrophoresis showed that the DNA was sheared down from a 10 kb chain into a slew of differently sized fragments ranging from 1 kb down to much less than 0.5 kb. This data indicates that the shearing protocol was very successful in producing fragments small enough to hopefully resist aggregation.
DNA Ladder
Gel electrophoresis. The first lane is the DNA ladder, the second lane is the unsheared DNA and the last lane is the sheared DNA.

Since I learned that new CTAB cannot be obtained until the middle of July, I am considering using negatively charged citrate gold nanoparticles and wrapping them in positively charged lysine molecules before mixing them with the newly sheared DNA.

Don't do drugs kids. Except caffeine... lots of caffeine.

Friday, June 17, 2016

Calf Thymus DNA

I started this week by cleaning and concentrating the nanoparticles that I made last week. On Wednesday I finally had my DNA to play with and I started by reconstituting it in TE buffer. The DNA I am using is a very common and cheap type of DNA extracted from a calf's thymus. It can be anywhere from 8-15 kb in length so its too long to make it through the centrifugal filters I have used previously for the PSS. I made 10 mg/mL stocks of the DNA and passed it through a syringe to hopefully shear it some and reduce the viscosity. After I reconstituted the DNA, I titrated into some of the gold nanoparticles at concentration ranging from 0.1 micrograms/mL to 1000 micrograms/mL and measured the UV-vis spectra at each concentration (see graph below).
We noticed that there was positive shift in wavelength with increasing DNA concentration which could be an indication of the DNA wrapping. We also saw some severe aggregation starting with 10 micrograms/mL that could be an indication of the formation of some sort of superstructure arrangement of the nanoparticles on the DNA. It will be possible to confirm if such a structure exists if we are able to visualise the nanoparticles on the TEM. However, the TEM is under maintenance right now so it is hard to now for sure. One superstructure that is possible and has been the subject of a lot of literature is a model histone (see picture below) which would have interesting implications.
I made up solutions of varying concentrations between 1 microgram/mL and 10 microgram/mL and measured their spectra to find a concentration that has DNA binding but no aggregation. The spectra showed the same increase in wavelength with increasing concentration but it also showed that the wavelength actually starts decreasing between 8 micrograms/mL and 10 micrograms/mL. It also shows that 2 micrograms/mL has a noticeable shift with little to no aggregation (See graph below).
Next week. I will attempt to clean off the excess DNA and measure both the bound and unbound DNA in the solutions.

June 17th - Making Nucleosomes

                This week I began the process of making nucleosomes. On Wednesday, we received a 500 mL supply of whole chicken blood, and I started the process of purifying 50 mL of it. Because the volume is “whole” chicken blood, it contains plasma, red blood cells, and white blood cells and platelets (see the image from the American Red Cross below). Birds are unique because their red blood cells contain nucleosomes, so the first step of the procedure was to isolate those cells. Such isolation was achieved by centrifuging the samples multiple times, and achieving a separation similar to the image below. The supernatant was the plasma, which was discarded, and the white-film on top of the red blood cells was also discarded.
                The next step in the procedure was to lyse the red blood cells, thus releasing the nuclei. Again, the samples were spun in the centrifuge and after successive spins, we were left with a clean, white precipitate. Over the next few days, we will work towards isolating just the nucleosomes from the nuclei.

Friday, June 10, 2016

Color Changing Kool-aid

Its the first week of research in Summer 2016! This summer I'm working on looking at how DNA interacts with CTAB gold nanoparticles while my friend Celina in Professor Thompson's lab continues the work with PSS coated nanoparticles that I started last summer. This week I mostly worked in Professor Thompson's lab changing the tubing on the nanoparticle flow-reactor and attempting to make a liter of CTAB nanoparticles using it.

Flow reactor used to make up to a liter of CTAB nanoparticles at a time.
It took a few tries to get the right reactant volumes for the monodisperse nanospheres we wanted. The first batch seemed, from its properties and UV-vis, to possible contain nanocubes. These nanocubes conveyed a really interesting property to the solution that made it a rusty red color in normal light and  a violet color when held up to sunlight.

In the end we managed to make nanoparticles with a smooth UV-vis around the right wavelength and normal DLS and Zeta measurements.

Next week, I am hopefully going to get some DNA and be able to start wrapping it around the nanoparticles.

First Post of 2016! June 10th, 2016

Summer research kicked-off this week for both Savannah and I, as we began to work on new experiments. I focused on two projects this week: analyzing data that Abby took during the summer of 2014, and doing preliminary reading on nucleosome core particles (NCPs). I will be working on both projects throughout the summer.
                The data Abby took in 2014 is very similar to the data I analyzed last summer, except there was ion competition between a monovalent and a trivalent cation charge neutralizing a hexagonal array of DNA (as opposed to divalent and trivalent). Abby had already computed the average concentrations of each element in the samples, and I was just compiling the data into graphs (see below). Problems with this such “raw analysis” of the data is evident though: the total cation-to-anion charge ratio easily exceeds one. This problem arises because the trivalent ion dissociates into a divalent ion in high chloride concentrations, which is not taken into account in the graphs below. But evening knowing the dissociation constant, simple chemistry cannot be done to determine the respective concentrations of the divalent and trivalent cations because trivalent ions favorably bind to DNA. Therefore, more theoretical work must be done to understand the system.
                Primarily throughout the summer, I will be working on the nucleosome core particle experiment. Last summer, Abby set the foundation by writing a procedure to make the NCPs, and doing some preliminary experiments with them. I plan to build off Abby’s work by improving the NCP procedure and doing more experiments with them. This week, I read and tried to understand her procedure, and hopefully next week I will start making the NCPs!