Tuesday, 1 August 2017


DAY 20: 24/07/17
The results from DAY 19's PCR is shown in figure 16.

It can be seen from figure 16 that the PCR with primers NRP1-K185-REV didn't work on DNAs #5-1, #5-2, #6-3, #6-4, even ad adapted conditions (lower annealing temperature, 48oC). So, a new PCR was prepared with NRP1-HBT-REV primer mix instead. Results are shown in figure 17.


It can be seen from figure 17 that there is a very light DNA band being emitted for #6-3 and #6-4. However, it is not very clear, so a new set of results were ran with the resting PCR. This time the exposure was changed in order to try and see the DNA bonds more clearly. Results are shown in figure 18.


It can be seen from figure 18 that there are DNA bands, however, the signal is weak (hence the need to increase exposure). Figure 19 is the same results as shown in figure 18, but with lowered exposure, showing no DNA bands at all to the naked eye (I thought this was a cool thing to show.. how looking at things from different 'angles' can change your interpretation of things).


From figure 15, it can be seen that N5-3 and N5-4 give a nice DNA signal, so those 2 PCR were sent to be sequenced against HBS2 primer.

DAY 21: 25/07/17
Today I will practice to do serial dilutions and spotting assays using a 96 well plate and a pinning tool (apparently this tool is very expensive, so I shall have to be extra careful with it)

Dr. MacNeill prepared beforehand 3 yeast cultures: wild type, N5-1 and N5-3. In order to do the dilutions, the OD600nm of each culture was measured, giving wild type OD = 0.26, N5-1 = 0.24, and N5-3 OD = 0.23. As it can be seen, the OD values for all 3 yeast cultures are of a close range, so there was no need to dilute the cells to OD600nm of 0.1 (also, this is just a practice day). Once the OD values were figured out, the cell dilutions was prepared, and cells were pinned to YES4 plates using the pinning tool (exciting bit). Dr MacNeill showed me step by step how do all of this, because I will have to do it on my own from no on.

In preparation for tomorrow, I prepared a culture for N5-1 and wild type (WT) pre-cultures (previously prepared by Dr. MacNeill), and left it to grow overnight at a shaking bath at 30oC.

DAY 22: 26/07/17
Today I did everything I learnt yesterday (on my own) using the N5-1 and WT cultures I prepared yesterday. The plates were incubated at 20oC, 32oC and 35oC to see how temperature influences growth.

The OD600nm values for the WT and N5-1 cultures were 0.17 and 0.19, respectively. Therefore, there wasn't a need to dilute the solutions to an OD value of 0.1.

Today pre-cultures for WT, N1, N3, N5, N8 and N10 DNAs were prepared and allowed to grow overnight at 32oC.

DAY 23: 27/07/17
Today I made cultures from the pre-cultures prepared yesterday, and the cultures were allowed to born overnight at a 30oC bath with shaking.

The plates prepared on DAY 21 and 22 were analysed. They still need to be left to grow for longer, however, it was possible to see the effects of the dilutions and how well presented a plate is by using a pinning tool (instead of manually pipetting all solutions into the agar plate). See this photo so you know what I mean:


DAY 24: 28/07/17
So today I used the WT, N1, N3, N5, N8, and N10 yeast cultures, prepared yesterday, to do serial dilutions and spotting assays.

The OD600nm values for the WT, N3 and N8 yeast cultures were below 0.1, so they were allowed to grow for ~30/40 minutes longer in the shaking water bath at 30oC. Apart from culture N10, all the other cultures, N1, N5 had their OD values of 0.11 and 0.13 (slightly above from 0.1, the desired OD value). All cultures were adjusted to have an OD value of 0.1, either by diluting or letting the cells grow more. Once all cultures had their OD value of 0.1, the serial dilutions were prepared

The serial dilution solutions were pinned on YE4S plates + HU (hydroxyurea) plates, with different HU concentrations: 0, 3, 6, 9 and 12 mM. Solutions were also pinned on YE4S plates to undergo UV light treatment. The UV light wavelengths were 50, 100, 150, 200 and 250nm. All plates were incubated at 32oC.

INFO: The purpose of growing cell at 'damaging' conditions (hydroxyure, UV light) is to see if the mutated cells (N1, N3, N5, N8, N10) are able to grow and if so, when they stop being able to grow.


DAY 16: 17/07/17
After a 2 weeks break, where I went to London to visit my friend, I'm back in the lab, and excited for the next part of my internship project.

Today I did miniprep DNA for colony PCR using N1, N3, N5, N8 and HBT plasmids.

INFO: what was previously called plasmids 1, 3, 5, 8, 10 and b3.4 are now called N1, N3, N5, N8, N10 and HBT, respectively.

The PCR was prepared in the following order: MW maker, N1, N3, N8, N10, N5 and HBT, where NSEQ1-RS1 primer mix was used for N1, N3, N8 and N10 DNAs; FS1-RS2 primer mix used for N5 DNA; and 5NSEQ-HBT-PCR-REV primer mix used for HBT DNA.

DAY 17: 18/07/17
The PCR prepared yesterday was ran on agarose gel to see the results. The results are shown in figure 10.
Figure 10: Agarose gel analysis of DNA MW marker against PCR solutions N1, N3, N8, N10, N5 and HBT, respectively

As it can be seen from figure 10, the PCR didn't really work for most of the DNA samples, since only the primers are giving out a signal for DNA samples N1, N3, N8, N10. However, for DNA sample N5, there are two visible strands, suggesting that PCR has fully worked, showing the prier strand and the DNA part of interest strand.

To check whether the DNA samples were ok, a new PCR was prepared. This time, using FS1-RS2 and NSEQ1-RS1 primer mixes. Results are shown below in figure 11.

Figure 11: Agarose gel analysis of DNA MW marker against PCR solutions N1, N3, N8, N10, N5, HBT (treated with FS1-RS2 primer mix), N1, N3, N8, N10, N5, HBT (treated with NSEQ1-RS1 primer mix), respectively.

As it can be seen from figure 11, the DNA samples are ok because, at least for FS1-RS2 primers, all DNA samples gave signal at the point of interest, suggesting the primers were the problem on the previous PCR reaction. Furthermore, it can be seen that the solutions treated with NSEQ1-RS1 primer mix didn't give signal for the DNA area of interest.

To check whether it was the primer not working, a new primer mix with NSEQ1-RS1 primers was prepared to do a new PCR.

DAY 18: 19/07/17
The results from yesterday's PCR is shown in figure 12.


It can be seen from figure 12 that the PCR didn't work, as only the primers themselves are emitting signal, thus, suggesting that the NSEQ1-RS1 primers combination is not binding properly to the DNA's part of interest.

In order to identify which primer is not working properly, new primer combinations were prepared and tested on N1 and N3 DNAs. The first new primer mix combination is NSEQ1-RS2 (RS2 have worked before with FS1 primer, as shown in figure 10 and 11). The second new primer mix combination is 5NSEQ-RS1. Results are shown in figure 13.


From figure 13, it can be seen that N1 DNA treated with NSEQ1-RS2 primers, show a very light DNA band. This suggests that some DNA from the part of interest is giving out signals. Furthermore, N3 DNA treated with NSEQ1-RS2 primers did not give any signal other than the primer itself. Neither N1 or N3 DNAs, treated with 5NSEQ-RS1 primers, gave any signals. These results suggest that the NSEQ1 primer is the problem in the NSEQ1-RS2 primer mix and we don't have enough data to determine which primer is the problem from the 5NSEQ-RS1 primer mix (results from figure 10, regarding the HBT treated with 5NSEQ/HBT-PCR-REV were inconclusive).

In other to obtain more data, other colonies of the HBT  tag colonies were treated with 5NSEQ-HBT0PCR-REV primer mix. Results are shown in figure 14.

INFO: The primer mix should only bind if the HBT tag is present in the DNA.

Figure 14: Agarose gel analysis of DNA MW marker against HBT tag colonies 3.1, 3.2, 3.4, 3.5, 3.6, treated with 5NSEQ-HBT-PCR-REV primer mix.

It can be seen from figure 14 that out of 5 HBT tag colonies, only one seems to actually have the HBT tag. The colony with a HBT tag seems to be colony #5-2.

DAY 19: 20/07/17
The only colony with a HBT tag seems to be colony #5-2, in figure 14, but since none of the others appear to have the HBT tag, which they should have (in theory), a new PCR with all #5 colonies and #6 colonies, as a control, was ran with different primers. The colonies were treated with NRP1-K185-REV primer mix, and also NRP3-NRP4 primer mix. There are two colonies for #5 and two for #6, so they were labelled #5-1, #5-2, #6-3, #6-4, and each colony was treated with each set of primer mix.

Furthermore, from figure 10, it can be seen that only N5 had a DNA signal, so DNA from all N5 and N6 colonies were prepared and treated with FS1-RS2 primer mix for PCR. There are four colonies within N5 and four within N6, so they were labeled N5-1, N5-2, N5-3, N5-4, N5-6, N6-6, N6-7 and N6-8.

The results for all #5, #6, N5 and N6 colonies are shown in figure 15.


From figure 15 results, it can be seen that the NRP1-K185-REV primer mix didn't work, as there is no DNA signal visible. However, that was expected because primer K185-REV optimal annealing temperature is lower than 60oC, so a new PCR with annealing temperature at 48oC was ran for #5-1, #5-2, #6-3 and #6-4 DNAs treated with NRP1-L185-REV.


DAY 13: 26/06/17
Today I will do colony PCR using 1, 5, 8 and 10 DNAs. This is to ensure DNA is going out signals. The gel was wrongly ran at 20V instead of 100V, but that's okay because the whole purpose of this PCR was to ensure that the DNA present is strong enough to give out a signal, and it is, as we can see in the image below.

DAY 14: 27/06/17
Today the only thing I had to do was to plate out the b3.4 1/100 dilution and 3 1/100 dilution grown solutions.

Plate out the solutions were fairly quick, so to do more things in the lab, I helped Dr. MacNeill with his own research buy doing colony PCR with some of his plasmids DNAs.

DAY 15: 28/06/17
Today PCR was ran using 1, 5, 8 and 10 plasmid DNAs with NRP1-NRP2 and NRP3-NRP4 mix primers. Results are shown in figure 7.


As we can see from figure 7, PCR results suggest that the PCR didn't work, so everything was repeated with optimising conditions, until it worked and the DNA gave out signals.

➢ A 'wildtype' was induced and a new primer replaced NRP2 was used: K185-REV primer (figure 8)
➢ Optimising conditions also involved changing the annealing temperature in the PCR reaction (annealing temperature went from 60oC to 48oC) (figure 9)

Results are shown in figure 8 and 9.


Ok, so Dr MacNeill is going on holidays with his wife, which means, I get some free time myself!!! 😚😃

Thursday, 27 July 2017


DAY 10: 19/06/17
The western blot prepared on DAY 9 was ran and scanned. Image is shown below:


The results didn't show a clear protein purification and that could be, for instance, while loading the gel, solutions ran from one well to another, mixing things up. However, it was unclear why the protein wasn't fully purified. Independently of the results not being clear, it was a good practice to make a western blot myself as I had never done it before (at uni, when we used one we were given a 'bought' ready one).

Today, spores solutions for b3.4 and 3 transformed plasmids were prepared and incubated overnight at 32oC

DAY 11: 20/06/17
Today, the spores solutions prepared yesterday, DAY 10, were plated out on EMM + Histidine + Adenine plates, and all 16 solutions (4 colonies from each plasmid) from plasmids 1, 5, 8 and 10 were plated on YE4S plates (YE4S medium had ampicillin and chloramphenicol added to it, in order to avoid bacterial contamination).

INFO: YE4S medium is a rich medium, and therefore all fungi should grown in it. EMM, however, as mentioned before, is a minimal medium and only the cells adapted to overcome the nutrients deprivation in the medium will survive.

So, for now any further step in my experiments require waiting for results, waiting for cells to grow. Thus, tomorrow will be a day off. Whoop, whoop I will sleep much much later than I'm used to these past weeks! 😎

DAY 12: 22/06/17
Today all plates were analysed to see colony growth. Also, because I've been using so many plates, I've prepared some more YE4S plates, and also YE4S + 5FOA plates for future use.

Again, in order to proceed with my experiments, waiting time to allow cells to grow more was required, so the weekend will be pro-longed with tomorrow! 😁


DAY 5: 12/06/17
Today's plan was to confirm whether plasmids 3 and 7, in fact, contained the insert at the correct orientation. This will be done by preparing solutions of plasmid 3 and 7, and N1, analysing them and sending them to be sequenced in a lab in Dundee (that's nice right, to have a lab that will do all the sequencing for you, and it's not even that far away..all you need to do is to send/receive a post).

Solutions of N1, 3 and 7 plasmids were prepared by mixing the DNAs with enzyme EcoRV, water and 10x cut smart buffer. A loading dye was added to each solution to help the solutions to 'fall' into the agarose gel wells. The solutions were loaded in an agarose gel and ran, the results are shown in figure 3.


Results from figure 3 just seems odd, and even the DNA MW marker that we saw last week looks odd compared to figure 1 and 2. So, since there was 'extra' of the N1, 3 and 7 plasmids solutions, another gel was ran. The results are shown in figure 4.


This set of results looks much better, however, it can be noticed that solution 3 wasn't ran properly as it seems that the gel was 'damaged', possibly by the pipette while loading solution 3. For that reason, from now on, only solutions 7 and N1 will be analysed. Note, however, that we can't just tell whether the primers are in the right direction only by looking at this figure. So, all solutions were sent to be sequence (in Dundee.. remember I mentioned?). Once they've been sequenced, their sequence can be compared to the 'expected sequence' (N1).

While waiting for the sequencing to be back, spores solutions (haploid cells) were made from the plates prepared on DAY 1. About 10 colonies from each plate was resuspended in water and helicase enzyme, then incubated overnight at 32C. 

INFO: why helicase enzyme you may ask? well, helicase digest diploid yeast cells, and also digests the cell walls of ascus cells (cells with 4 spores present), releasing haploid cells (spores). Why are the ascus cells you may ask? Well, that's because the spores were grown on EMM. EMM plates are poor plates, where only yeast cells capable of producing all amino acids are able to grow. Also, at starving conditions, diploid yeast cels will undergo meiosis, forming ascus cells. Sometimes, the cell wall naturally breaks, releasing haploid cells.

MORE INFO: In day 1, 4 plasmids were transformed, as shown below:
1. leu1-32/leu1-32
2. Ura4-D18/Ura4-D18
3. his3-377/his3-377
4. ade6-M210/ade6-M216  
⇨ The yeast strand used in day 1 is sp280 cdc27+/cdc27::ura4+ (cdc27+/''cdc27Δ), where :: means replaced with.
⇨ It can be seen that leucine, uracil, histidine and adenine are unable to be made due to their gene's alleles mutation/deletion. So, how are these yeasts going to grow? Well..

  •   Plasmid 1 is fixed with Leu2 plasmid (leu2 ≣ leu1+), where ≣ means equivalent to. 
  •  Plasmid 2 is fixed with ura4+ in cdc27 
  •  Plasmid 3 is fixed by the addition of histidine in EMM 
  •  Plasmid 4 doesn't need to be fixed when diploid. This is because Ade6 is a dimeric protein, and as you can notice, different alleles of the same chromosome were deleted. Thus, ade+ is produced. However, in haploid cells, adenine is added to EMM to enable the yeast to obtain adenine and grow. 

DAY 6: 13/06/17
Today the spores solutions prepared yesterday, on DAY 5, will be placed on EMM + Histidine + Adenine plates. Histidine? the cell is unable to produce it, and adenine? spores are haploid, and therefore, an adenine- mutation on plasmid pHBLA-Cdc27-N8, deletion of a gene will stop adenine production (in diploid cells, due to being diploid, the dimeric protein having two different alleles of the same chromosome being deleted would still allow the production of adenine+). So in order for the cells to be able to grown in EMM plates in haploid form, both histidine and adenine are added to EMM. Each spore solution was plated as it is, but also diluted in 1/10, 1/100 dilutions.

So this all so far have been part of my project, but because my part of internship now requires a lot of waiting (e.g. wait for cells to grow). So I helped Dr. MacNeill with one of his projects by preparing plasmid solutions for 3 plasmids,  Z1, Z2 and Z8. Then prepared restrictive enzyme solutions, with EcoRV and HinIII enzymes, to run an agarose gel. This was good laboratory practice, as I had learn how to do this, so now I'm having the chance to emphasise my learning and put it in practice.

Two gels were ran because for the first one, a mistake occurred (yes I know... tragic!!). So, I mistook the loading dye (which helps solution to sink in the gel wells) with MyTaqRed (an element for PCR reactions). In my defence, they both had the same colours 😝. So I added MyTaqRed instead of loading dye, and when I realised my mistake (I realised although the liquid looks the same colour, pink/purple, the tubes are different... the MyTaqRed is not labeled and the loading dye is labelled), I added loading dye as well.. I knew that some ingredients that make up MyTaqRed wouldn't have affected the reaction, but I was afraid the TaqPolymerase enzyme would, so I repeated everything again. The Results are shown in figure 5 and 6. Figure 5 represents the first reaction with the addition of MyTaqRed and figure 6 represents the repeated version, without the MyTaqRed.



Figure 5 shows a better gel than figure 6. This is because, after all, the MyTaqRed, did not affect the results, but also it looks like the mixtures were better mixed, and the pipetting better carried out. This is concluded due to, in figure 6, the Z1 HindIII restrictive enzyme solution shows an extra band that shouldn't be there, suggesting that the enzyme didn't cut the plasmid properly. Also, the Z2 EcoRV restrictive enzyme solution shows that the gel was slightly damaged. Apart from that both gels look clear. Furthermore, to analyse the results, the bands sizes were compared to the DNA plasmids sequence, confirming that the plasmids solutions were correct. Sequences were analysed using Enzyme X program software.

DAY 7: 14/06/17
Today the sequences from the DNA plasmids 3 and 7 were analysed, to confirm whether they had the right insert at the right orientation. Also, plasmid b3.4 was transformed into diploid strain: pHBLA-Cdc27-Ne (with K187R mutation)

The sequences for DNA plasmids 3 and 7 were analysed using first '4 peaks' software, to analyse the quality of the peaks, and check where about the sequence start/stop to be clear. The ALIGN website was used to analyse the DNA plasmids sequence 3 and 7. Both sequences were compared to the expected sequences for a plasmid with an insertion in the right orientation, reverse orientation and a plasmid without the HBT insertion. After analysing the sequences, it was concluded that both plasmids (3 and 7) had the HBT insert at the right orientation (both identity and similarity shows 100% for the correct orientation.

Plasmid b3.4 was transformed into diploid strain: pHBLA-Cdc27-Ne (with K187R mutation), and transformed cells were plates and incubated at 32oC. This transformed plasmid is the plasmid that I didn't have on DAY 1 (2nd Uni binding site).

DAY 8: 15/06/17
Today we analysed under a light microscope how the spore plates prepared in DAY 6 and also the transformed plasmic pHBLA-Cdc27-N3 prepared in DAY 7, as well as plasmid 3.

INFO: Although the plates from DAY 6 still need to be left to grow for longer, it can be seen that the diluted plates show less and more spread out spores (haploid cells). It can also be seen that some plates show bacterial contamination, but we will worry about that later on, when the yeast colonies have had time to grow more.

Today a SDS gel was also prepared to be used tomorrow.

DAY 9: 16/06/17
Today I started the preparation of the SDS gel running/western blot. The protocol is long and even tho there is only two line written so far, it took me the whole day to complete it. The blot itself was allowed to dry to be used on Monday again.

Sunday, 11 June 2017


DAY 1: 6/6/17 
I was super nervous for my first day and I didn't know what to really expect! Once I arrived, I was greeted by Dr. MacNeill, and we went through the plan for the week. He showed me around and where everything was in the lab. My first task was to read all the healthy and safety rules on my cubicle (yes, I got a whole cubicle desk for myself.. how cool is that?! super cool I say!). Once I read everything, I was ready to head to the lab, and feel like a true scientist.

Dr. MacNeill had prepared beforehand 4 plasmids (he attempted to make 5, but for some unknown reason, the 5th plasmid just wasn't having any of it, and decided not to work). Today in the lab, I attempted to mutate each binding site found in the cdc27 (p66 in humans) subunit. Plasmid 1 had its phosphorylation sites mutated. Plasmid 2 had one of its ubiquitylation sites mutated. Plasmid 3 had its sumoylation (SUMO) site mutated. Each plasmid was transformed into cdc27/cdc27del yeast diploid cells and incubated at 32oC for about a week (to allow cells to grow and colonies to form).

DAY 2: 7/6/17
Today's task was to cut, purify and ligate DNAs (plasmid sample and insert DNA), then transform E. coli. The plasmid used as N11 plasmid, and it was digested with NotI-HB restriction enzyme. The inner DNA cdc27 (spcdc27-HBT) alleles were also digested with NotI-HB restriction enzyme. Before ligating both the plasmid sample and the insert DNA together, their solutions were run in an agarose gel, against a DNA molecular weight tag, for comparison (see figure 1). Once ligation was done, a 50ul and a 150ul solutions were placed in LB+ ampicillin plates and insulated overnight, allowing transformation to occur.
NOTES: bacteria was used for transformation because it grows quicker than yeast.

From figure 1, it can be seen that the plasmid is A and that the insert DNA is B. This can be told, not only by being organised and labelling the solutions being placed in the agarose gel, but also, because of the molecular weight of both solutions. The insert DNA is smaller than the plasmid, which means it will travel further in the agarose gel.

DAY 3: 8/6/17
Today's task was to screen colonies by colony PCR and set up culture for mini-preps. However, one of the enzymes required for colony PCR hadn't arrived yet (it was still in Germany.. so we waited). However, I still did some lab work: the plasmids prepared on DAY 2 were grown, and 8 colonies were placed into 8x tubes containing LB Broth and ampicillin, and incubated in a shaker overnight to form bacterial culture.

NOTES: two primers will bind to the transformed plasmid, however, a signal will only be transmitted if both primers are pointed to each other (correct orientation). If it occurs that more than one insert DNA is present in the plasmid, then the shortest sequence will be amplified (correct orientation). The longer sequences can happen to be amplified, but they will mostly likely serve as a template for the amplification of the shortest sequence. Furthermore, the longer sequences are hard to be identified in an agarose gel.

DAY 4: 9/6/17
Today, the primers arrived, so it was possible to carry on colony PCR using the bacterial culture prepared on DAY 3. All the 8 PCR solutions were allowed to undergo 25 cycles in a PCR machine (took ~1 hour 15 mins), then solutions were run in an agarose gel against a DNA MW solution, for comparison. Solutions were placed in the gel in the order: DNA MW, 1, 2, 3, 4, 5, 6, 7, 8 (see figure 2 for results).

From figure 2, it can be seen that plasmid 3 and 7 have the strongest signal, suggesting that those two plasmids underwent transformation and ended up with the right sequence at the correct orientations. So, due to the results from figure 2, it was decided to prepare/transform plasmids 3 and 7 into cdc27/cdc27del diploid following the same protocol as in DAY 1.


This summer, I have secured an internship for the Summer of 2017 in the School of Biology at the University of St Andrews, under the supervision of Dr Stuart MacNeill. This internship opportunity is supper exciting because I will obtain hands-on experience of state-of-the-art laboratory techniques! Also Dr MacNeill has  been successful in his career. He has contributed with ~ 75 publications and is a SULSA (Scottish Universities Life Sciences Alliance) Reader in Translational Biology. Dr. MacNeill’s research is focused on chromosomal DNA replication and genome stability, and I'm really hoping to learn as much as possible from him. 

My internship will focus on chromosomal DNA replication, more specifically, in cdc27 subunit of DNA polymerase. I will try and keep you guys updated on what is happening inside the lab! Also If you're interested in extra readings, there is a few papers listed in the references section. And if you want learn a little bit more about my internship aims and why what I'm doing is relevant, please read the background information section, shown below. 

Background Information

Successful chromosomal DNA replication is essential for maintaining genome integrity in all forms of cellular life. Defective DNA replication can have a wide variety of effects on genome structure and information content, such as sequence deletion, insertion and duplication, point mutation and chromosome fusion. In mammals, defects in replication can lead to developmental defects, growth impairment and tumour formation.
The proposed work centres on two family B DNA polymerase enzymes with distinct roles in eukaryotic chromosome replication: DNA polymerase δ (Pol δ) and DNA polymerase ζ (Pol ζ). At the replication fork, Pol δ is essential for lagging strand DNA synthesis and may play a role in leading strand synthesis also (1). Pol δ also acts in a variety of DNA repair pathways and several recent studies have shown that mutations in human Pol δ bring about tumour formation. Pol ζ is translesion synthesis (TLS) polymerase (2). This enzyme replicates through sites of damaged DNA but does so with low fidelity. Pol ζ activity therefore contributes to cell survival following DNA damage, by ensuring that the chromosomes are fully replicated, but at the expense of introducing mutagenic changes into the DNA. Gaining a complete picture of the structure, function and regulation of these enzymes is crucial to fully understanding their cellular roles and the broader contributions that they make to maintaining genome integrity.
At the molecular level, human Pol δ comprises four subunits: PolD1, PolD2, PolD3 and PolD4. PolD1 (also known as p125) is the catalytic subunit, possessing polymerase and proofreading exonuclease activities, whereas PolD2 (p50), PolD3 (p66) and PolD4 (p12) are accessory subunits whose contributions to Pol δ activity remain largely unknown (3). It was recently shown that PolD2 and PolD3 are also subunits of human Pol ζ, which is a heterotetramer of PolZ1 (catalytic subunit Rev3), PolZ2 (Rev7), PolD2 (p50) and PolD3 (p66) (2).
Fission yeast is an excellent model for investigating the function of these enzymes. Like its human counterpart, fission yeast Pol δ is heterotetrameric, comprising orthologues of PolD1 (Pol3), PolD2 (Cdc1), PolD3 (Cdc27) and PolD4 (Cdm1) (4). This is not the case for the more widely studied budding yeast Pol δ enzyme, which lacks PolD4. Fission yeast Pol ζ has not been characterised biochemically but is likely composed of Rev3, Rev7, Cdc1 and Cdc27.
The proposed project focuses on Pol δ/Pol ζ shared subunit Cdc27, the fission yeast orthologue of human PolD3 (p66), and its potential regulation by post-translation modification (PTM). Cdc27 is a 372 amino acid protein. The first 160 amino acids bind to Cdc1, the fission yeast PolD2 protein, while interaction sites for Pol α–primase and the sliding clamp processivity factor PCNA are found in the C- terminal region of protein (spanning residues 293-332 and 362-369, respectively) (5,6). By comparison with human PolD3 (p66) (7), the N-terminal region likely forms a winged helix-turn-helix (wHTH) domain. In contrast, the C-terminal region may be largely unstructured.
Recent proteome-wide analysis has identified two ubiquitylation (Lys185, Lys187) (8), one sumoylation (Lys317) (9) and five phosphorylation (Ser160, Thr161, Ser163, Ser167, Thr187) (10,11) sites within the C-terminal domain of Cdc27 protein. The sumoylation site maps within the minimal Pol α–primase interaction motif mentioned above (6), suggesting that sumoylation may function to modulate this interaction. None of these PTMs is essential for Pol δ function, as we have previously shown that cells expressing a truncated protein lacking amino acids 161-372 are viable (12). Aside from this, the functions of the PTMs have not been investigated.

The overarching aim of the proposed work is to gain insight into the potential roles of ubiquitylation and sumoylation in modulating PolD3 function. Using fission yeast as a model system, the student will use a combination of biochemical and molecular genetic methods to investigate the role(s) of these PTMs. Through this work, we hope to provide evidence that the identified modifications play a role in modulating the properties of Cdc27, thereby opening up new avenues of research into the regulation of Pol δ and Pol ζ activity in yeast and ultimately, human cells. 

1. Lujan, S.A. et al. (2016) Trends Cell Biol, 26, 640-654.
2. Makarova, A.V. and Burgers, P.M. (2015) DNA Repair (Amst), 29, 47-55.
3. Tahirov, T.H. (2012) Subcell Biochem, 62, 217-236.
4.   Zuo, S. et al. (1997) Proc Natl Acad Sci U S A, 94, 11244-11249.
5.  Gray, F.C. et al. (2004) BMC Mol Biol, 5, 21.
6.  Reynolds, N. et al. (2000) EMBO J, 19, 1108-1118.
7. Baranovskiy, A.G. et al. (2008) Cell Cycle, 7, 3026-3036.
8. Beckley, J.R. et al. (2015) Mol Cell Proteomics, 14, 3132-3141.
9. Kohler, J.B. et al. (2015) Nat Commun, 6, 8827.
10. Koch, A. et al. (2011) Sci Signal, 4, rs6.
11. Kettenbach, A.N. et al. (2015) Mol Cell Proteomics, 14, 1275-1287.
12. Tanaka, H. et al. (2004) Nucleic Acids Res, 32, 6367-6377. 


DAY 20: 24/07/17 The results from DAY 19's PCR is shown in figure 16.                                       It can be seen from figu...