While we've had a few nice days recently, it's been pretty rainy over the last few weeks.
Finals are almost over and the students are moving back home. It's kind of a bittersweet time of the semester. The weather and feel of campus brings back memories of just a year ago when I was finishing up my undergraduate career. After being at Clark for 5 year (9 semesters and 2 summers), I'm really going to miss it. I know I'm not looking forward to packing up and moving home myself, but I have plenty to do before then.
I'm defending my Master's thesis on May 22nd, and I hope to be completely done a few days before the end of the month. I have a first draft of my thesis completed, which will probably head over to my thesis committee once my adviser okays it. I've already made corrections from a working (almost complete) draft, which was nearly sufficient for submission anyway. I'm pretty excited about this. Once I submit the draft to the other two members of my committee, I'll focus on finishing my powerpoint presentation.
My powerpoint presentation for my defense will probably be about 45 minutes long. Just thinking about it right now is making me nervous. However, I hope to have a final draft of my presentation completed more than a week before I defend, giving me ample time to practice. This will be the third presentation I've given this semester, and by far the longest, most difficult, and most important. While I'm nervous now and I'll be nervous on my defense day, I just know I'll knock my presentation dead. The closed question period may prove to be much more difficult, but I know I've done more than enough work to complete my Master's.
Between now and the time I move out, I'm almost working on some diatom cultures. I hope to have a large array of diatoms lined up for my professor to work with this summer. Below I discuss part of this process:
And, as a bonus, I made this video in my spare time–how to tie a bowtie!
I've received my new primers and already run a couple different reactions with some results that prove to be promising and others that are frustrating.
Yeah, encouraging :]
The first thing to do with a new set of primers is to run them in a normal PCR reaction to see if the primers amplify the correct length of DNA. While I could do this with my super concentrated, ultra clean plasmid DNA that I used to transform my diatoms (which would serve as the ultimate positive control), I decided to be be bold and try to amplify my complimentary DNA (cDNA) samples. This cDNA was made from mRNA that was extracted from diatom cells exposed to different environmental conditions. Because the mRNA we're looking for contains GFP, anything that is amplified essentially means that our plasmid DNA that we transformed into this line of diatoms is being expressed, which is a great, great thing.
So I ran this PCR reaction with my new primers to amplify my cDNA and this is the gel I got:
There are some pretty convincing bands in that gel which is really encouraging. It appears that all but two of my reactions (6 out of 8) gave us at least some PCR product.
Nicccccce :D
I then ran a real-time PCR reaction called a standard curve, where the source DNA is serially diluted ten fold (I ran 1:1 through 1:10,000 dilutions). For this reaction, I do go ahead and use the plasmid DNA control to give us the cleanest results possible. This reaction allows us to see how efficient the primers are at doubling the amount of DNA product at each PCR cycle.
You can see in this gel (in the first 5 lanes) that a lot of PCR product is being produced. Each reaction hit their saturation point which is why each reaction looks the same even though they had drastically different amounts of starting DNA template. The great this about real-time PCR is that we can see on the computer screen how each reaction was amplified in real time, and see where each reaction it a ceiling amount of DNA. In this reaction I could see that even the 1:10,000 dilution easily hit this ceiling before the 40th (and final) cycle.
With these two encouraging results--the proper amplification using my new primers in a standard PCR reaction and strong amplification in my standard curve using plasmid DNA--I went ahead and tried amplifying my cDNA in a real-time PCR reaction.
Unfortunately, I ran into the same problem I've been having for a few weeks now (see the bottom half of this post).
Sad face real-time PCR :[
Not all of my reactions amplified, and those that did didn't amplify as cleanly as I wanted them to. I ran a gel of my second real-time PCR reaction, which visualizes the two amplified reactions (lanes 3 and 6 starting from the top).
While this is certainly a set back, I have a couple of things I'm going to try. Most importantly, I've ordered a new kit to run the real-time PCR reactions, since the kit I was using was "old." Next, there are a few things I can do to ensure my reactions are as balanced and clean as possible. Finally, I'm going to drop the annealing temperature of my real-time PCR. While I was using the same annealing temperature from my standard PCR, there are two main things that differ between my standard and real-time PCR reactions. First, the salt concentrations are most likely different (although that's a bit annoying to look up, but it's on my to do list), which I found out while screening my diatoms can really screw up a PCR reaction. Second, I used the mysterious "Q-solution" provided by the company Qiagen in their PCR kit when setting up my standard PCR reactions. This may also have significantly changed my standard PCR reaction. While the real-time PCR reaction really should be working with my current set up, it's very possible that my primers are finding it difficult to "seek out" and bind to the sparse cDNA that I want them to amplify.
Last post I talked about amplifying DNA by real time-PCR, which measures the number of amplified copies at the end of each cycle, giving researchers "real time" numbers of DNA copies. To do this however, you need appropriate primers to get the job done. Not only do they have to be specific enough to work only for the DNA you want amplified, but I'm learning there are other tricks you need to abide by.
Let's back up a little bit though and look at transcription and translation. After all, I'm after the mRNA transcripts that are made in this process. With my genetically engineered construct, protein synthesis starts when transcription factors bind within the cloned 5' untranslated region (UTR) and begin transcription at the promoter, transcribing all of the way through the eGFP open reading frame (ORF) and through the 3' UTR. Now we have an mRNA transcript with part of the 5' and 3' UTR intact at either end of the eGFP coding region. This will serve as the template for translation, which begins at the start codon of the ORF and ends at the stop codon. The 5' and 3' UTR are not translated, hence their UTR moniker.
My project is aiming to measure the amount of mRNA transcript in cells under different environmental conditions. It's not easy to measure mRNA by itself, but it is very easy to measure DNA. Using mRNA as a template, you can make complimentary DNA using the enzyme reverse transcriptase.
Reverse transcriptase starts at the 3' end on an mRNA molecule and transcribes a complimentary strand backwards along the mRNA. However, reverse transcriptase will eventually fall off (represented by the fading orange triangle), so smaller mRNA transcripts work the best.
If I have this mRNA transcript that I want to measure through RT-PCR (after I've converted it into cDNA with reverse transcriptase), I need a primer to amplify the eGFP coding region.
I first made this primer pair to test for the presence of eGFP in my diatom cell lines. It amplifies most of the eGFP coding region by attaching at points just inside of the gene, as you can see below.
Unfortunately, these primers aren't working very well when it comes to applying them to real time-PCR.
My control real time-PCR reactions have worked pretty well for me each and every trial I've run. The control reactions use primers to amplify the endogenous genes we're manipulating in our system, which serve as a good comparison to the experimental reactions.
You can see on the graph at right that the primers amplifying endogenous genes work pretty well, developing curves within an appropriate cycle range (the number of cycles until a noticeable amount of product can be measured).
However, when I look at the graph for the transgenic lines using the above primers to amplify eGFP, I get a graph like this on the left. The amplification lines are severely delayed and do not approach the same level of product by the end of the reaction.
In addition to amplification plots, the real-time PCR application on the testing computer also shows graphs that display the melting point of the double stranded DNA molecules. These graphs can be very informative when troubleshooting real time-PCR reactions.
Here is the dissociation curve of the endogenous amplifications:
Oh boy, it's that curve crisp and clean.
Here is the dissociation curve of the transgenic amplifications:
Yeah, not so much. This curve is extremely messy and non uniform. This provides further evidence that my primers for this reaction might not being working as well as they should be.
After talking to my adviser and seeking some advice online, I've found a couple of parameters to follow to make better primers.
First and foremost (and going back to my bit about reverse transcriptase starting at the 3' end of the mRNA transcripts), my adviser let me in on a secret: I should be using primers that amplify near the 3' end of the transcript since that will be the highest quality region of cDNA as it is transcribed. Using this knowledge, I am working on primers that amplify in the area represented by the orange square, just outside of the 3' UTR. I also tweaked the settings which the primers conform to, based on information I found on other university websites. Yay for Google and other scientists!
The past few days in lab I've been working on these primers and planning out my semester of science ahead of me. Just this week I put together my thesis committee (Justin Thackeray who I had for genetics three years ago, and David Hibbett, my undergraduate adviser) and I've begun re-reading some primary literature and will soon begin reading more broadly and in depth in preparation of writing my thesis. I've also started putting together bits of my paper, which I should have done a while ago.
Anyway, that's all for now, really.
If you've gotten this far, watch my latest YouTubes video regarding this topic matter:
In this video I discuss designing a colony screen protocol for my transformed lines of diatoms. In one of my recent posts, I posted this picture diagram which oversimplified the process:
While the PCR steps are the same (starting in the top left panel with the diatoms being combined with the PCR mix), I discovered that diatoms grown in liquid culture (i.e. their natural state) needed extra care in preparing them for a colony screen.
Let's step back a bit and quickly talk about colony screens (in case you didn't read my above Tumblr post). A colony screen is a modified PCR reaction*, which is a cyclical amplification of a short sequence of DNA, exponentially copying the targeted DNA strand. (*Even though PCR stands for polymerase chain reaction, PCR is usually said out loud in conversation as PCR reaction.) Often the DNA source for a PCR reaction is a purified, such as plasmid DNA purified by a process to separate plasmid DNA from proteins and genomic DNA.
But colony screens use a colony of cells to supply the DNA for the PCR reaction. By initially lysing the cell by cooking it at a high temperature for a period of time, the cell's DNA is released into the PCR reaction mix and then amplified in the reaction. Colony screens are used to screen cells for the presence of DNA--that is if the targeted sequence of DNA is present, it will be amplified by the reaction. If a reaction gives a positive band on a gel (bottom right of the colony screen chart), then we know that the DNA from the cells in that particular reaction also have the DNA of interest.
This colony screen method should in theory work for other cells, like diatoms. My initial trial successfully amplified GFP from diatom colonies growing on agar plates, but it didn't work for diatom cultures growing in liquid media, a distinction I didn't make at the time.
I soon realized after a failed trial of screening diatoms solely from liquid cultures that the residual salt water from their growth media was throwing off the delicate chemistry of the PCR reaction. This graphic paired with the above video discusses just this.
Since this video was recorded, I've made further steps in my project. In order to test cultures of diatoms sooner for their presence of GFP and at a much higher volume, I've been designing gene specific primers to amplify regions of DNA in the diatoms, that would yield bands of DNA if the GFP is present. There are four main sets of primers I've been designing and will use in PCR experiments:
By using the standard NR-eGFP-NR construct as an example, the sets of primers would like the above picture, where orange highlights show where primers would amplify if the sequence exists in the DNA. Primer (1) would yield segment [a]: a portion of the 5' UTR, eGFP (enhanced GFP), and a portion of the 3' UTR. Primer (2) would yield segment [b], a portion of the 5' UTR and a portion of eGFP; primer (3) would yield a segment [c], portion of eGFP and a portion of the 3' UTR. Primer pairs to amplify the (1, 2, & 3) regions will be made for all of the gene constructs, that is NR-eGFP-NR, NR-eGFP-Actin, NiR-eGFP-NiR, and NiR-eGFP-Actin. Therefore only a specific set of primers will be used per diatom culture, depending on which plasmid it is supposed to have.
All of the plasmid however will be amplified with the fourth and final primer pair. Primer (4) would yield an eGFP segment [d] if the diatom cells had it. This will be the most important primer result out of all of them.
Once we have these primers at hand, I can screen many colonies at once. All I need to do is take a small DNA sample from a colony, add it to a PCR reaction, and run the reaction. I should be able to quickly decipher which colonies have GFP based on an electrophoretic gel.
And now I can talk about the next step of my project: the transformation of diatoms with the plasmids I've created.
Now I am undergoing the process of selecting lines of transformed diatoms for the next step in our project. But we've recently realized that the light intensity in our growth chamber is much brighter than previously published experiments growing transformed cultures of diatoms. Increasing light intensity reduces the amount of chlorophyll made by plants and algae. Why is this important? Two reasons:
Less chlorophyll will reduce the pigmentation of our cells, making them harder to see once we transfer them to liquid culture.
Our gene giving resistance to our antibiotic (that allows us to select for positive transformants) is driven by a chlorophyll-associated protein, which means expression may be reduced in higher light intensities. If chlorophyll expression is reduced by high light levels, then this may reduce the activity of the chlorophyll-associated protein that drives the antibiotic resistance in our diatoms, which means less resistance to the antibiotic in the media. This ultimately means reduced or no growth.
Now, our diatom cultures have been growing okay on their agar plates, but I'm beginning to wonder if they're growing slower in liquid cultures, where they're more likely to absorb more light. To combat this, I'm growing some liquid cultures behind layers of porch screen to reduce light levels penetrating the liquid cultures. In the below picture, you can see the different diatom cultures I have growing right now:
Transformed diatoms grow on agar plates (left foreground) an in liquid media (left background and right).
I've been growing diatoms in 3 mL cultures in little dishes, which you can see in the back left. Groups of 3 dishes sit in a petri dish to allow easier transportation. These dishes however require 3 mL of culture just to cover the entire bottom of the dish, which means the cells are going to be fairly diluted. I also started 1 mL cultures in the test tubes on the right, where porch screen blocks out a lot of the incoming light. Some of the 3 mL cultures also have screen on top of them to block out some of the light.
Aside from trying to grow our cells in liquid culture, I've been looking at them under a fluorescent microscope to try to determine whether our cells are fluorescing due to GFP expression or whether it's residual glow from chlorophyll. UV light from the microscope excites all pigments in the sample, and then I look at the sample through different filters that only allow certain wavelengths of light to show through. However, our current filter set up hasn't allowed a clear delineation between GFP and chlorophyll yet. Most of our pictures look something like this:
...but they usually have a lot more background color in there too.
My next steps will be to either design or order commercial GFP GSPs--gene specific primers for GFP. That way we can run a PCR on some diatom DNA and determine whether they have the GFP gene they are supposed to be transformed with. Because I have a great control (the original plasmid DNA), I can test a whole bunch of diatom colonies at once and be able to select appropriate lines of diatoms faster.
I've been having trouble transforming E. coli cells with my plasmid vector, within which is a small DNA fragment (my NiR terminator) that I will want to restriction cut out. By transforming bacteria with the plasmid vector, I'll make additional copies of the plasmid and be able to freeze and save the plasmid for later use if necessary.
My lab mates and I spent several weeks trying to figure out why our transformations were doing so poorly and why we were receiving such low plasmid yields from transformed bacteria. I myself figured out that one problem was the ampicillin used to make the agar plates upon which we grow our bacteria had degraded over time, and that the antibiotic was not selecting strongly enough to weed out bacteria with plasmids and bacteria without plasmids. This is the reason why we were not getting good plasmid yields and another reason why our bacteria were not growing when transferred from "old" plates to new agar plates with freshly made ampicillin.
We also concluded that the bacteria cells we were transforming were not up to par to yield the results we needed, so we ordered some new transformation kits.
But in order to successfully clone PCR product into a plasmid vector to transform into bacteria, the PCR product needs to be freshly made. In order to get new PCR product, I re-amplified older PCR product in the same reaction I ran before. I ran four different reactions using the PCR DNA in four different DNA concentrations: 1:1, 1:10, 1:100, & 1:1,000 (lanes 2, 3, 4 & 5 in the picture below respectively). This way I can determine which reaction had too much starting DNA and too little. After my reaction, I ran part of it on a gel to see how each reaction went. I definitely got much larger yields in the 1:1 & 1:10 dilutions (there was probably too much DNA even), so I used the second dilution (1:100, lane 4) to clone into the plasmid vector.
I used PCR product from lane 4 to clone into a vector plasmid for the transformation.
Using the vector plasmid, I transformed them into the bacteria and let them grow over night on an agar plate. I then performed a colony screen, which is a PCR reaction using single bacteria colonies to supply the DNA. That PCR reaction yielded the below gel:
While this is a slightly messy colony screen gel, several of these colonies should suffice!
What we're seeing in this gel is the molecular ladder at the top and then 10 different colony screen reactions. They're pretty streaky, which is probably because there was a lot of bacterial DNA in each PCR reaction. What I wanted was a single band at around 700 basepairs, which is roughly half way between the two second most right bands on the ladder. As such, lanes 4, 6, 7 & 8 are good candidates for colonies that have my plasmid with the correct insert.
Sometimes science (my project) isn't linear, and I've been getting caught up in this recently with my posting.
I wanted to do a series of posts and videos on the process of transformation and the completion of my first NiR plasmid, but what I found was that sometimes things don't work out the way you want.
So, let me backtrack a bit.
As I mentioned before, it appears I have transformed bacteria colonies that have my PCR insert. Great! But I've been having trouble getting the insert to amplify out of the plasmid once again. I grew up several bacteria colonies that looked like they had my insert (white colonies on X-gal) and performed a plasmid prep that yielded very little plasmid DNA. I need a decent amount of this plasmid to allow me to digest (cut) out and obtain the insert.
Now, I'm trying to do yet another PCR reaction in a much larger volume (50µl rather than 10-20µl) using the plasmids I obtained from my lame plasmid prep. If this works, I'll have a lot of copies of the NiR terminator insert, which I can then slice off the ends with restriction enzymes. Then, the insert would be ready for the next step.
Last time I checked in, my PCR reactions that were supposed to add restriction sites to either end of the nitrite reductase (NiR) terminator region appeared to work and work well. When I ran the PCR reactions on a gel, the amplified DNA bands were really strong and were the correct length.
This gave me the go ahead to continue the path in isolating & altering the NiR terminator in order to yield a sequence of DNA to fit the final NiR plasmid for my project's experiments.
Now I need to insert the NiR terminator into a vector plasmid, transform it into bacteria, and digest the NiR terminator back out of the plasmid to double check that the terminator I got on the gel from my PCR reaction (above) is the correct piece of DNA before I insert it into the NiR plasmid to complete the final NiR plasmid.
We use bacteria to amplify pieces of DNA because of their quick generation times. If you insert a plasmid into bacteria, they will duplicate the plasmid as if it were their own DNA as they grow and divide. A plasmid is a ring of DNA, which is essential for this to work, because bacteria will cut up and destroy any loose pieces of linear DNA. In order to get our NiR terminator to be duplicated by the bacteria, we insert it into a vector plasmid first. The vector plasmid is designed to accept small pieces of DNA from PCR reactions, lock in that piece of DNA within the plasmid. This plasmid can then be transformed into bacteria.
Bacterial transformation is really easy. Once the vector plasmid complete with our PCR DNA is ready, we add the plasmids to specially-altered E. coli cells, incubate the cells on ice for a short time (to lull them into a false sense of security), and then transfer them to a hot water bath (42°C) for thirty seconds. Thirty seconds is all we need for the bacteria cells to panic and scream "WHAT IS HAPPENING TO ME?" This prompts the bacteria, because they are stressed, to take up any DNA in their environment. Well good thing the only DNA in their environment is the plasmid we gave them! Through the heat shock, a large amount of bacteria should have taken up our plasmid. We then grow the bacteria over night while they recuperate, divide exponentially, and make copies of our plasmid. This process is summarized by the cartoon below:
Three PCR reactions, three beautiful bands. Looks like digesting the insert and then amplifying it really did the trick. This means I can use my PCR reaction to transform some bacteria!
The idea is to insert our PCR reaction into a plasmid vector, which we then transform into E. coli. We do this via heat shock, which causes the bacteria cells to take up plasmids. We then will grow the E. coli, conduct a plasmid prep to collect all of the plasmids they grew, and then I will perform another digest to recut out the terminator region (the PCR product). This way, I can be sure the restriction sites were correctly added onto the terminator region, which I need in order to insert it into the rest of the plasmid I already have.
I'm transforming the cells as I write this, and will be back soon to give updates. Yay science!
Today I'm running a digest on a plasmid in an attempt to cut out a portion of DNA.
This plasmid is the nitrite reductase (NiR) gene terminator region cloned into a vector (called pCR4). Just outside of where the terminator region should be inserted is a series of primer points and restriction sites. EcoRI restriction sites flank just outside the cloning site, and as such I'm running an EcoRI restriction digest. The digest only takes an hour at 37°C on a heating block (right), and is a simple reaction containing just restriction enzymes, buffer, DNA, and water.
I took a quarter of the restriction digest reaction (5 of the 20µl), added 1µl loading dye, and ran the reaction on a 2% agarose gel:
Here we can see in the second lane my digest. The really bright band is the vector (most of the DNA in the plasmid) and the smaller band (further along on the gel) in my insert, just under 1,000 base pairs. The ladder (the top and 5th lanes) is a 1kb ladder--moving right to left, the smallest band is 500bp, then 1kb, 1.5kb, 2kb, 3kb, and 4kb, etc. I like to use a 1kb ladder most of the time because it's really easy to use, and I can quickly tell which band is which because the 3kb mark is the brightest in the ladder. in my gel here, the vector sequence is larger than 3kb, and roughly equal to the 4kb band (which was expected). The insert that was digested out in the reaction is just short of 1kb (which was also expected.
I then ran the rest of the reaction on a second gel (which was a sort of a waste, but I always want to check my reaction before I look to do anything else with it), because my digest worked. I wanted to cut out the smaller band and use that in a PCR reaction, to make it easier for the primers to amplify them. Since I've been having trouble getting the primers to work on this plasmid, we decided we might as well try this. So I ran the rest of the digest reaction in lanes 2 and 3--you can see where I cut out the bands, which I did with a razor blade (which is the black silhouette on the right).
I then melted the agarose gel that I cut out with my DNA band, and cleaned up the solution. By binding the DNA to a small filter, I could clean the DNA and remove the gel. I then yielded (what I hope is) the digested plasmid insert. I'll use this cleaned up insert from the gel in a PCR reaction overnight tonight and hopefully get some better results.
Below is a cartoon representing the NiR sequence I'm working with. For now, I'm just focusing on the terminator region, which is the 3' untranslated region (UTR) sequence just after the stop codon. By sequencing the DNA we're working with or looking up the desired DNA sequence online at a databank website, we can model the DNA sequence and figure out where the start and stop codons most likely exist. We can then make primers to amplify specific regions along that DNA.
The blue rectangle represents the NiR terminator that we're amplifying: it's just a little bit longer on either side of the actual 3' UTR, which means we're sure to amplify the entire 3' UTR.
I've already talked about how I've made an inducible expression plasmid to test the mRNA stability of nitrate reductase (NR) transcripts in vivo in the diatom Thalassiosira pseudonana. The plasmids run by using a promoter and terminator sequence to run the expression of a reporter gene (we're usingGFP). We have a set of NR plasmids, one with a terminal region of NR and one with a terminal region of action (which has nothing to do with nitrogen assimilation). The former plasmid should mimic endogenous activity, whereas the latter plasmid should not mimic the normal conditions in the cell. NR reduces nitrate (NO3-) to nitrite (NO2-), which is then reduced to ammonium (NH4+) by nitrite reductase (NiR). As such, in the scheme of nitrogen assimilation in diatoms, it makes sense to test the regulation of NiR as well.
To do this, I've started work on a set of NiR plasmids. They will also run the inducible expression of GFP, but with their own promoter and terminator regions. Both will have their NiR promoter region, whereas one will have the NiR terminator and one will have the actin terminator region. Both sets of plasmids will be transformed through particle bombardment and then in vivo expression can be measured (through GFP activity).
As it stands, we have a plasmid that contains the NiR promoter & GFP but not the proper terminator. The NiR terminator was cloned into a separate plasmid as part of the process in amplifying out the terminator region of interest. Currently, I'm trying to amplify the terminator out of the plasmid, and in doing so add restriction sites to the plasmid.
The cartoon below represents the series of steps that we have to do in order to manufacture the desired fragment of DNA with restriction sites at the beginning and end of it, which allow us to easily cut out the DNA fragment and place it into a plasmid. Right now, the NiR terminator is represented by the red box. It was amplified out of the entire NiR gene to yield a small piece (between 500bp and 1kb) of the the gene. This piece was at the very end of the open reading frame and extended past the 3' UTR (the terminal region). This was done in a PCR reaction, and the PCR product was inserted into a vector, transformed into bacteria, grown, and then the plasmids were isolated once again the yield the below plasmid.
What I want to do is amplify a smaller portion of the insert out of the plasmid, and make a bunch of copies of it through a PCR reaction. A second set of primers (the green lines) will amplify within the region of the insert, while adding restriction sites. Currently, I've having problems getting this PCR to work because less than half of the primer fits to the DNA of the current insert, while the other half is going to add the restriction site. However, I changed the protocol for the PCR reaction I'm running at the moment. I changed the annealing temperature for the first 10 rounds of my PCR and then I'll bring it back up to what I ran it yesterday (a PCR reaction that did not work, lanes 3-5; right--below right my PCR samples loaded; the faint blue samples are the ladders I used, lanes 1, 2 & 8; the red samples are my PCR reactions that used Coral Load, a special PCR buffer/loading dye combination, lanes 3-7).
...NiR terminator amplification to be continued...
