Tag Archives: intro

The SciFund Experiment: Yeast Adaptation to D2O

It has taken me quite a while, but I’m finally at the point where I can begin the experiments that my SciFund quest funded. To refresh your memory, here is my project proposal:

Water is arguably the most important molecule in the universe. It’s a simple molecule that is composed of two hydrogen atoms and one oxygen. But did you know there are different types of water molecules? Every element has alternate forms known as isotopes, and hydrogen is no different. A common isotope of hydrogen, known as deuterium (D) which is twice the mass of hydrogren, can bond with oxygen to make heavy water (D2O).

Immediately after heavy water was first purified from naturally occurring water in the early 1930s, it was discovered that most organisms cannot survive in pure heavy water. It was also shown that increased (but not toxic) levels of heavy water significantly affect many systems in these same organisms, like fertility, metabolism, temperature regulation, and many more, all of which are essential for healthy organism function. Interestingly, on a cellular level the increased mass of heavy water may affect chemical processes. Not many studies have been performed in this area, unfortunately, because many experimenters ignore the effects of water even though it is by far the most abundant molecule in these experiments.

Because of the presence of deuterium in naturally occurring water, life may have evolved essential uses for deuterium. I plan to study the effects of heavy water on E. coli and S. cerevisiae (baker’s yeast). I will be growing cultures of these microbes in water with varying amounts of heavy water (from 0% to 99.9%) and comparing characteristics between the cultures, looking for effects in growth, development, appearance, and other physical differences.

But that isn’t specific enough to describe what I actually intend to do with the money. The full explanation of the experiment is way cooler!

Initially I’m going to continue the experiments from April and May that compared the growth of yeast grown in DI water, DDW, and D2O. Each time I did a time trial experiment I would start from scratch to show that yeast grows much slower in D2O. This time I’m going to carry over the yeast each day. Over time I hope to show that the yeast is adapting to growing in D2O and begins to develop at a similar rate to how it develops in DI water and DDW.

Once that happens I’ll be doing two experiments.

First I’ll be looking for phenotype differences between yeast grown in H2O vs yeast grown in D2O. I’ll be looking for shape, size, motility, etc of the yeast using the lab’s light microscopes.

In the second experiment, I’ll flip the script on the yeast. Once I get the yeast to adapt to D2O, I would like to determine if H2O is as harmful to D2O adapted yeast as D2O is to H2O yeast (aka natural yeast). I’ll do this by growing the D2O adapted yeast in DDW/DI water and recording the growth. No matter what happens with this experiment, the results will be interesting.

3-piece ligation: BpALS, EpBR, 5′-bio/int-bio adapter

A few days ago I did a 3-piece ligation that didn’t turn out so well. Today’s ligation is more like the traditional ligation that I was supposed to do with the exception that I’m using the pALS anchor instead of the pRL anchor.

I mentioned earlier, that BpALS (pALS anchor digested with BstXI) is better overall because it is longer and makes the optical tweezer operations a little easier and better.

pBR322 can be digested with EarI or SapI for use in our construct for DNA unzipping. I haven’t done any studies that show which method produces better results in terms of final yield, but I’m pretty convinced SapI will because you only have to perform one gel extraction. If you digest pBR with EarI you get two linear fragments. And according to separate analysis by Koch and myself the overhang that we need for the unzipping construct ligation is the longer piece (in the gel image linked above it is the brighter top band).

The benefit to following this method is you can perform a three piece ligation (essentially ligating the entire construct together in one shot) because the EpBR piece (pBR322 digested with EarI) can only ligate to the adapter, and the BpALS/BpRL piece can only ligate to the other end of the adapter (both adapters that I’ve been using work the same way).

And so today I’m doing the proper three-piece ligation reaction. The reaction setup and method can be seen in the table below:

pALS PCR digestion with BstXI

The pALS anchor is special in that it has two purposes. The first is that it is immediately usable in DNA stretching experiments because it has a dig molecule (to stick to glass) on one end and a biotin (to stick to microspheres) on the other. The second is that I can digest it with BstXI restriction enzyme and use it as the anchor segment for unzipping DNA.

And when it all works well, pALS is much more useful than the pRL574 anchor (1.1kb). It’s extra length makes it easier to calibrate the optical tweezers for unzipping and we can get higher forces in the optical trap by using bigger beads. (Note: The tweezers are the entire device, and the trap is the focal point of the laser in the microscope. So the trap is a subset of the tweezers.)

With the huge success of the pALS PCR yesterday, I’m going to digest some of it and then ligate this piece to EpBR and both adapters. But first here is my digestion reaction:

Annealing the adapter oligos

I purchased some oligos and now I need to create the adapter duplex from those oligos. This requires an annealing reaction which is super simple to do. Basically you just mix your oligos together, heat them to 95C and then slowly cool the mixture. Nature takes care of all the leg work. I’ve done this reaction 3 different ways:

  1. Heat a cup of water to boiling in a microwave, then remove the water, place your annealing mix in the water (make sure the top is floating), and allow the water to cool on a lab top. Basically allow the water to cool to room temperature (RT).
  2. Put your mix into a heating block and heat that to 95C (or close to boiling). Once the mix has been heated, remove the block from the heating unit and place on a lab top to cool to RT.
  3. Put your annealing mix in a PCR machine which can control the temperature very specifically. Create a program that will: (1) heat the mix to 95C for about 5 minutes, (2) slowly lower the temperature to about RT or 4C or whatever cool temp you want, (3) hold at that low temp until you are ready to remove it and move on.

For today’s reaction I will be using option 3. My protocol for the experiment is below. Unfortunately there is no easy way to verify the annealing reaction is successful. Well that’s not 100% true. You can run an SDS-PAGE gel which has the ability to resolve very short DNA sequences, but I don’t really have the equipment for that right now. I do have a high resolution gel that I’ve been wanting to try. Hmmmm…

And here is a link to what my annealing buffer is: Annealing Buffer Recipe.

Shotgun DNA Mapping: Microspheres

Today I need to buy microspheres. In the first experiments we used spheres with a diameter of 0.5um (or 500nm), and over time we eventually switched to using 1.0um beads. The reason is because: 1) big beads are easier to see if they are tethered or not, 2) you get better tweezer forces with the larger beads, and 3) the big beads clump less. The only drawback to using the larger beads is that there is a lot of repulsion between the beads and our glass surface so our DNA tethers need to be longer. I invented the 4kb pALS anchor to solve this very problem.

Here are some pictures of the different sized beads in the tethering environment:

And for completeness here are some old notebook entries regarding those pictures:

And here is a video that shows the tethering results of the DNA experiments listed above:

We want beads that are coated in streptavidin (or some form of avidin) because this molecule creates a very strong bond with biotin which is attached to our DNA for stretching/unzipping experiments.Most commonly, you can order beads with streptavidin, but some companies offer alternatives like avidin or neutravidin. In my experience neither works any better or worse than streptavidin. And in the case of these experiments the bond either holds or it doesn’t.

But believe it or not, it is hard to quantify the effectiveness of the beads. As you can see in the pictures above, both bead concentration and DNA concentration can affect tethering efficiency. And I have suspicions that the sonication process (what we do to prevent the beads from clumping) may affect the streptavidin in some way: in my head the vibration shakes off the molecules from the beads.

With all that said, there are places that I trust buying beads from. In the past I’ve purchased beads from Bangs Labs, Invitrogen, and Poly Sciences. I’ve never noticed that any bead from any company seems to work better than the rest. Because of this I think I’ll order a new stock of beads from Bangs because they pretty much only make beads (so they should do it the best). Note: I just remembered that Bangs, and Poly Sciences may be the same company and it turns out they are affiliated in some way. So I suppose there really is no difference between the two.

Update: I’m placing my order with Bangs Labs. I’m ordering 0.53um beads and 1.04um beads, both coated in streptavidin, and neither are fluorescent.

Shotgun DNA Mapping: Unzippable DNA

Unzipping the complete construct…

We’ve discussed the anchor DNA and the adapter duplex, but we wouldn’t be able to measure unzipping forces and shotgun DNA mapping would fail if we didn’t have any DNA to unzip. As I’ve mentioned several times in this series of posts, the entire construct is assembled through a reaction known as ligation.

For the purposes of this experiment, the reaction works as follows: an enzyme known as DNA ligase looks for compatible ends of DNA and attaches them together. In our construct those ends are the overhangs that I referred to in the other posts. And the construct is designed so that the anchor can only attach to one end of the adapter and the unzipping segment can only attach to the other end of the adapter.

Now technically we can use any piece of DNA to unzip. The catch is that we need to use a plasmid to get the overhang that we need. As I said earlier, one side of the adapter can ligate to the anchor. The other end’s overhang is created by a cut from the enzyme EarI and is specific to the plasmid pBR322 and any other plasmids that have the same multiple cloning site. For instance, for shotgun clones (which will be explained much later) we use pBluescript II, and the enzyme SapI cuts the plasmid with the exact same overhang as EarI does in pBR322.

Because of the proximity of the SapI site to the multiple cloning site, we can stick any piece of DNA into the plasmid for cloning. Then we can cut the plasmid with the unzipping insert with SapI and then ligate this long piece to our unzipping construct.

In calibration experiments, we use pBR322 to test to make sure we have unzipping. And eventually we will move up to use the pBluescript clones that I made a few years ago. Although I have a feeling I’ll be doing that all over again.

Shotgun DNA Mapping: The Unzipping Adapter

Ignoring the circle, the adapter duplex (the middle piece, red) will be the topic of today’s discussion.

The ligation reaction that I keep referring to requires three pieces of DNA. They get fused together all in one shot, that is slightly complicated. The most crucial of which is the adapter duplex, because without it the anchor and the unzipping DNA would not attach and the reaction would yield nothing. And because of how important the adapter is, this has been the source of my troubles for the past 4 years. But before I go into that, let me tell you about the duplex.

It’s called an adapter duplex because it is actually two single stranded pieces of DNA. We call them the top and bottom strand. They are short DNA sequences manufactured from biotech companies. In the past we’ve used Alpha DNA, but I’m thinking of trying someone new. How short are the strands? The bottom strand has about 35 bases and the top is only a few bases longer. Compare that to the anchor sequence which is either 1100 bp (base pairs) or 4400 bp or the unzipping sequence which can be as long or as short as we want (but typically around 3000bp for calibration sequences).

Once our single stranded sequences arrive via mail (we call these short sequence oligonucleotides, or oligos for short), we need to bind the top and bottom strands together in a process called annealing. Most molecular biological reactions involve some kind of enzyme to help the reaction, but annealing is quite a natural process. DNA naturally wants the bases to bind to complementary bases (A-T, G-C) and even in single stranded form, the DNA will self anneal, that is bind to itself. So to get our top and bottom strands to stick together we just put them together in the same tube, heat it up to near boiling temperatures, and slowly bring the temp down so that the top and bottom strands find each other and bind. Once it’s cooled, the adapter duplex is formed and will stay that way unless heated to very high temperatures (near boiling).

There are three key features of the adapter duplex: (1) a biotin molecule, (2) a gap in the DNA backbone, and (3) two non-palindromic overhangs. The overhangs are designed to bind with a very specific sequence. One side can only bind with the overhang I mentioned in the anchor DNA, the other side can only bind with the overhang contained in the unzipping DNA. Right now that particular sequence is very specific to cutting plasmid pBR322 with the enzyme SapI (and any other plasmids that share similar properties).

The biotin is necessary for unzipping. The biotin has a high affinity for streptavidin which coats the microspheres we use for optical tweezing. Typically the biotin in our bottom adapter strand is near the start, but not at the start of the sequence. In more recent iterations, we moved it to the 5′ end completely or added a poly-A overhang with several biotin there. The reason for this is because we’ve been having issues actually unzipping, which I’ll explain in another post. The hope was that by moving the biotin we would get better tethering efficiency and better unzipping. We ended up not getting unzipping results and the tethering efficiency studies were inconclusive.

See wikipedia, DNA article

The bottom strand has both the biotin and the gap (key feature 2), which actually plays a role in the unzipping. Since the tweezers will pull on this side, the gap was designed to aid in the unzipping. Basically the gap was the weakest point in the complete DNA chain and since the microsphere is so close to it the DNA would begin to unzip from this location. The gap is actually a missing phosphorus (the yellow in the image to the right), which prevents the anchor and the bottom adapter strand from connecting to each other.  In later iterations we completely removed the first base to make the gap wider, and the poly-A tail I mentioned was also used to prevent there from being any attachment.

Ultimately I never got unzipping to work. Oddly enough, I ran experiments that verified the ligation reaction worked, but could never get the completed structure to unzip. That’s what this new set of experiments is going to attempt. But before I get to that, I need to tell you about the unzipping DNA portion!

Shotgun DNA Mapping: The DNA Anchor

The complete unzipping structure being unzipped.

In order to unzip DNA, I need to create three pieces of DNA that I will then attach to each other through a ligation reaction. The first piece that I will discuss is the anchor DNA.

The anchor DNA is a very versatile piece of double stranded DNA (dsDNA). From this singular piece, we can choose to unzip DNA or stretch it because of a special sequence contained in the DNA near one end. I’ll get into this a little bit later. But first a couple of questions:

  1. Why is it called anchor DNA? The reason is because we use this piece of DNA to attach our entire structure to a glass surface. This is the point that anchors our DNA while we pull on it for either stretching or unzipping experiments. One of the bases is designed with a digoxigenin molecule attached to it and that base is placed right at the start of the sequence. In our tethering experiments, we coat our glass with an antibody for digoxigenin (dig for short), cleverly named anti-dig, and chemistry causes the anti-dig to bind with dig. You can understand a lot about antigen-antibody interactions here.
  2. How can we decide between stretching and unzipping? Because of how we designed the anchor DNA, we can stretch the anchor segment by default. That means once I produce anchor DNA I can tether it and begin stretching experiments. If we want to unzip DNA, then I take the anchor DNA and cut the end off (the side opposite the dig molecule) in a digestion reaction (more on this another time). That reaction gives me a small overhang (when one side of the DNA is longer than the other). From there I can perform a series of reactions that create the DNA sequence necessary to perform unzipping experiments. Notice that the anchor end is left unchanged, and that is what enables us to perform both stretching and unzipping experiments from this one piece of DNA.

Now the third question is, How do you make the anchor sequence? For this we need to know several sequences, possibly perform some cloning, and perform a reaction known as polymerase chain reaction, or PCR.

I’m not going to go into the details of what PCR is and how it works (google searching will reveal a lot more useful information than what I’d be willing to put here), but what I will say is that PCR allows me to make millions/billions of copies of a sequence of DNA starting with just a few strands of the original sequence and some short pieces of DNA called primers.

Our original sequence comes from plasmids. For the anchor sequence I have two possible starting points: pRL574 is a plasmid that dates back to Koch’s graduate days, and about a year and a half ago I created a brand new plasmid called pALS. Both plasmids are viable options, but serve slightly difference purposes:

  • pRL574 – for this plasmid we have several different sets of primers that allow us to make anchors of different lengths ~1.1kb and ~4.4kb. The 4.4kb sequence we use primarily for stretching experiments, while the 1.1kb sequence is used in unzipping experiments.
  • pALS – this plasmid only produces one length which is about 4kb. But this plasmid allows us to both unzip and stretch as I described above. It also has a couple of very unique features. First, if I cut it in the right spot, I can ligate the plasmid to itself through a special adapter sequence (to be described later). Second, it contains a sequence that is recognize by nucleosomes, that we could use for more complicated experiments down the road.

So as you can tell, I have some options available to me. Normally I would just pick one plasmid to work with, but I want to work with both and figure out which may be the more viable option down the road. In my next post, I’ll link to and list the sequences needed to make the anchor construct, with some explanations as to what everything is.

The Library of Congress: Science at Risk meeting

Back at the end of March, I was invited to a special meeting hosted by the Library of Congress. The meeting is entitled “Science at Risk: Toward a National Strategy for Preserving Online Science.” The meeting commences next week on June 26 and 27, and I am excited!

There are little details that I know about the event, but here is some info that I was forwarded:

Scholarly discourse, including interaction between scientists and the public, is rapidly changing and the ephemeral nature of this discussion on the web leaves it at substantial risk of being lost. Science blogs, the work of citizen scientists, and novel online publications like video journals are becoming the primary sources for understanding science in our times. These resources are almost exclusively online and increasingly at risk. The goal of this meeting is to begin identifying content that is valuable and at risk and to articulate next steps to ensure that this content is not lost.

In the face of this challenge, the Library of Congress, with generous support from the Alfred P. Sloan Foundation, wishes to join with other organizations to develop a national strategy for collecting and preserving science and science discourse which exists only in digital form on the open web.  The Library would welcome your participation in the invitation-only workshop “Science at Risk: Toward a National Strategy for Preserving Online Science” to be held over one and a half days on June 26-27, 2012.  The event will bring together scientists and representatives from online science projects, archivists, and the historians and other scholars who will increasingly depend upon the historical record.

From what I’ve seen the guest list is full of impressive people, and for me to be included among them is a huge honor.

I’m going to be speaking on behalf of open notebook science and scientists. I’ll present various open notebooks, including my own, and current methods for successful and useful open notebook science. As the week goes on I’ll have more details about this and I’ll be notebooking my preparations. Next week I’ll do my best to document the meeting.

If you are an open notebook scientist and would like your notebook mentioned/featured then feel free to contact me on twitter (@thescienceofant), email (anthonysalvagnoatgmaildotcom), in the comments below, regular mail, on facebook, or smoke signals (but please be in the ABQ area otherwise I may not see it).

 

Proof of Principle for Shotgun DNA Mapping (Redux)

You might have noticed all the recent activity with lack of context, well it’s Spring Break here at UNM and Steve and I are going into overdrive to finalize some papers that were left in preprint limbo. We have three projects to work on of various priority: (1) Shotgun DNA Mapping (what this article is about), (2) a paper that models Kinesin motility, and (3) a paper about the Repeating Crumley experiment that will be self-published via Google Docs.

So all the stuff that has been appearing in my notebook from both myself and Steve is all about the Shotgun DNA Mapping (SDM) paper. Our goal is to complete the paper, with some extra experiments that show how useful the SDM software is, and complete it by the end of the week (I’m guessing by Sunday because that’s when my parents arrive).

Right now we are refamiliarizing ourselves with the software. The original code and resulting paper were written in September of 2008 (yikes I’ve been in grad school for a while). The code was originally written by Larry Herskowitz and back then he had talents as a programmer, but he had no talents as an organized human being. So getting familiar with his mindset and looking for important programs and files is no easy task. There may be a lot we need to do and that’s what we are trying to figure out. Hopefully tomorrow we’ll be able to move on from this step.

But what code am I talking about? You probably thought I was just an experimentalist. Well you’d be mostly correct, but I have dabbled in programming some.

The code in question is the heart of Shotgun DNA Mapping. The SDM project was a two step experiment:

  1. Generate clones of random yeast genomic DNA sequences and unzip those sequences using our optical tweezers.
  2. Compare the force curves from the unzipped DNA to a library of simulated force vs extension curves. The library is generated from the yeast genome.

The paper that we are working on was a proof of principle for step 2. The results from step 1 would be published once we had working tweezers and unzippable DNA. Unfortunately we couldn’t get unzippable DNA, but maybe this summer I’ll be able to try again. In this paper we discuss how the simulation software works, how we match genetic information, and we present results using some old data Steve had from grad school.

Aside: It just occurred to me that this could be a success of open science if other groups had DNA unzipping data that they shared online, but alas the rest of the world is closed 🙁

Let’s discuss the software briefly so there is some context as to what Steve and I may write here for the rest of the week, and so you can understand the basic premise of the paper.

To get a brief understanding of how the tweezers work and how we are able to unzip DNA check out my intro to SDM here.

Now that you understand all that stuff let’s get into the software:

  1. The first step to SDM is to create a library of unzipping force vs extension curves for the yeast genome. We chose yeast because it’s DNA is bundled as chromatin which could be used for the next level of SDM, Shotgun Chromatin Mapping (SCM), and because our collaborators are expert yeast geneticists.
    1. We downloaded the yeast genome sequence from yeastgenome.org (back in 2008) and did a simulated restriction digest. By this I mean we looked for the XhoI recognition sequence (CTCGAG) in the yeast genome. From each recognition site, we created “fragments” that are 2000bp in length to use for the unzipping simulation.
    2. The unzipping simulation used a very simple equation to calculate the energy contained in a double stranded DNA sequence (dsDNA). The hamiltonian (as is  the term for energy terms) contained the energy of the freely-jointed chain (which is a model that describes a chain of paperclips, google it) and the base-pairing energy (ie A-T/G-C bond energies). That’s it! Remarkably that worked exceptionally well. The reasoning is that when you are unzipping you have two sequences of single-stranded DNA held together by the unzipped DNA (base-paired bonds).
    3. After the energies contained in an unzipping sequence is calculated we needed to extract force information. We did this by solving an integral of x(F’)dF’ numerically, where x(F’) is the extension depending on the freely jointed chain model. I don’t expect that to mean too much right now and I may have incorrectly explained it myself, but this will be made more clear later.
  2. Once we have simulated unzipping data we can begin to match data to the library we just created.
    1. I don’t understand the mathematics behind the matching algorithm all that much myself (right now), but from what I understand the matching uses a normalization that is routed in the difference between a polyA strand and a polyG strand (polyA has forces of ~9pN and polyG has forces of ~19pN, everything else is in between).
    2. Because of the nature of the simulation, at low extensions there is a lot of unreadable data, so we use a window between bases from 1200 and 1700 (we call this number index) to get better analysis. We also chose a 500 base window size above index 1000 because of the unzipping data that we were analyzing.
    3. The algorithm generates a score based on the difference between real and simulated force profiles in the window size stated above. In our system great matches are close to 0 and bad matches are close to 1.
  3. In graduate school, Steve had unzipped a plasmid known as pBR322 (purchasable from NEB). Using the software Larry had simulated the force profile for this sequence and hid the data in the library of yeast genomic simulated data. The matching algorithm managed to match the real unzipping data to the simulated data every time.

Like I said a lot, I don’t expect all of this to make sense, but as we rewrite the paper I’ll add thoughts about the project to the notebook. On top of that I’m sure we’ll have a lot of supplemental information to add to the paper via the notebook (so we can just site it). So while all this is new and confusing (if it isn’t then that’s awesome) be aware that the confusion will subside by the end of the week. We have a lot of exciting things coming in the next few days and you’ll all be the third to know about it (after Steve and I of course).