We also had presentations today. We updated the class on the addition of the caffeine and bleomycin groups and went into more detail about our experimental design. This presentation can be found in resources. We also did more research and reformulated our hypothesis on the relationship between the removal IRA2 gene and the resulting sensitivity to chemicals and toxins of the mutant yeast. After further research, we decided to use the chemical caffeine and the toxin bleomycin and test their effect on the growth of our mutated yeast. Bleomycin is a toxic drug used in chemotherapy. It works by causing double or single strand breaks in actively dividing cells in order to kill them and halt rapid cell growth and division. We hope that if we use the correct concentration in our agar plates, the bleomycin will limit the yeast cell's rapid growth without killing them all off. With the bleomycin and the caffeine, we will test their effect on the mutant yeast's growth by mixing it into the agar plates and comparing it to yeast grown on normal agar. In order to do this, we had to figure out how much caffeine and bleomycin to mix in with our agar. After some research, we determined that the amounts for 100 mL solid YPD media should be 7.7 uM caffeine and 3.2 uM bleomycin.
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Given this updated information, we finalized the number of replicates we would be working with. We decided to do the control group and three experimental groups each with the wild type strain and the IRA2 mutant strain. The control group would be grown in normal conditions on both agar plates and YPD medium at 30 degrees Celcius for 48 hours. The second group will be to test heat resistance and will be grown on agar plates incubated at 37 degrees Celcius for 48 hours. 37 degrees Celcius is signficant because it is the average temperature within the human body and also because it is typically too hot for successful yeast growth. The third group would be grown on agar plates with agar mixed with 7.7 uM of caffeine and the fourth group would be grown on agar plates with agar mixed with 3.2 uM of bleomycin. All of our groups will be repeated in a second trial, resulting in 16 agar plates and 2 viles of liquid YPD media.
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Using the plates we poured last class, we then prepared 8 of our 16 agar plates with both strains of yeast. We used the micropipette to put 10 uL of yeast onto the appropriate plates. We tried our best to pipette the yeast into a semi circular blob since our plan was to measure the colonies dimensions and determine their area in order to observe differences in growth. At the end of the lab, the plates were taken to the incubators, remembering that the heat resistance group would be grown in the 37 degree incubator. Next class, we will make the caffeine and bleomycin plates.
VIRTUAL LAB NOTEBOOK
Day 1- 8/24/16
Today we researched several different nonessential genes that are commonly found in yeast, including IRA2, TEP1, SAK1, TOS3, SDH4, and FUM1. After some thorough research, we chose the gene IRA2 as our group's gene to research more in depth. We learned about the gene and how its presence affects yeast's ability to grow in different conditions. We mainly researched what would occur if we removed IRA2 from yeast. We discovered that if IRA2 was removed, yeast would be able to grow at higher temperatures due to increased thermotolerance. We also discovered that IRA2 helps encodes proteins that regulate yeast's growth the amount of chitin made in the yeast cells. Chitin allows budding sites to form, so if IRA2 is removed, more chitin will result in more budding. The lack of IRA2 will also cause yeast to lose its ability to sporulate, or produce sexually. Yeast without IRA2 will also be sensitive to certain chemicals. For example, caffeine would change its color to brown and calcofluor white would change its color to white. We can use that sensitivity to see if our experiment was successful. Based on our research, we came up with a hypothesis stating that if the gene IRA2 is removed from yeast through genetic engineering, the yeast cultivated will be able to grow under high temperatures and produce more haploid cells than diploid cells that have increased sensitivity to certain chemicals.
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We presented these ideas to our class in a presentation that can be found in resources.
Day 2 - 8/29/16
Today we learned about double strand breaks in DNA and how a cell responds to a double strand break through homologous repair. We then brainstormed how to use that reaction to perform genetic engineering on yeast.
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Double strand breaks occur in cells DNA and trigger an emergency response. The cell launches into homologous recombination repair. First, specific enzymes called apurinic/apyrimidinic (AP) endonucleases go to the break and start to chew away at one strand of the DNA on both sides of the double strand break. This creates two pieces of DNA with long single strand tails at one end. Next, a homologous piece of DNA is located and unzipped by enzymes. Then, one of the single strands binds to the unzipped homologous DNA. The end of the single strand serves as a primer that attracts the DNA polymerase enzyme to the site. The DNA polymerase then adds nucleotides and transcribes the DNA. As this occurs, the new base pairs are separated from the single strand from the homologous DNA which goes back to its other strand. The homologous DNA only served as a template for a single strand of homologous DNA. After the DNA polymerase finishes transcribing the new single strand of DNA, it can then bind once again to the other single strand that was created by the AP endonuclases, therefore fixing the double strand break.
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Based on what we learned about homologous repair of double strand breaks, we hypothesized that we can use polymerase chain reaction, or PCR, a method of amplifying DNA, in order to create many copies of the yeast DNA. We then came up with a plan to use template DNA along with primers that we created online from the genome of yeast to inject the yeast cell with homologous DNA.
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In order to make the primers, we found the genome of yeast on an online genome database. We then copied that genome into a primer generator online. This website provided us the base pairs for a forward and reverse primer that would give us about 500 base pairs in each direction.
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Our primers were:
Forward primer:
AAC CCA AAG TAG CTC CTC AAA AAG GGT CAC
Reverse primer:
ACT TTG GGT GGG ATA ACA TCA ATG CTT CGAG
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The next question we had was how will we make the double strand breaks occur where we want them to? After PCR, we will have a large amount of DNA to work with during homologous recombination. This will ensure that a double strand break will occur at least once in the middle of the IRA2 gene, where we want it to occur. However, breaks will occur in other locations too. In order to let us know where the breaks in the IRA2 gene are, we can insert a gene for an antibiotic resistence gene where the IRA2 gene would normally be, in between the two primers. There is a gap here because the primers will ensure that the DNA is chewed away on both sides in order to create a hole in the middle where we can insert anything we want. We will insert an antibiotic resistance gene so that when homologous repair is occuring, the broken DNA will use the homologous DNA that we inject rather than other homologous DNA that may be in the cell already. Therefore, the antibiotic resistance gene will ensure that our DNA will survive exposure to bacteria and other DNA in the cell will be killed off, forcing the cell to use the homologous DNA that we feed to it in its repair of a double strand break. We are selecting against the homologous DNA that already exists in the cell so that the broken DNA is forced to use the DNA that we give it in order to perform a repair of the double strand break.
Day 3 - 8/31/16
On Day 3, we first started with practicing our micro-pipetting skills.
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Using a protocol given to us by our teacher (found in resources), we prepared a solution for polymerase chain reaction or PCR.
We figured out the amounts of DNA template, primers, IRA2, DNA polymerase, water, a buffer and dNTPs we needed for the PCR. For the water, buffer, dNTPs, primers and DNA polymerase, our teacher gave us a protocol that included the amounts for a 50 microliter reaction. For the template DNA, the amount for each group was variable based on which gene they were using. For us, IRA2's concentration was 31.8 nanograms per microliter. The final concentration of template DNA had to be under 250 nanograms so we did a calculation to determine that we needed 7 microliters of template DNA.
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We then realized that some of the amounts in microliters were smaller than the micropipette could pipette. We decided as a large class to combine some of the ingredients in a collective vile between all 4 groups, mix it up, and then extract 1/4 of the mixture for our own individual viles. We could only do this with the ingredients that everyone needed the same concentration of. These were the buffer, the dNTPs and the DNA polymerase. The primers, amount of water and amount of template DNA differed for each group. The group vile was made, mixed and kept on ice. Each group then took 1/4 of the mixture and mixed it with the appropriate amounts of their primers, template DNA and water. We then mixed it in a centrifuge machine to make sure it was well mixed.
In PCR, our template DNA was a yeast knock out (YKO) deletion collection strain with the G418 antibiotic resistance gene inserted in the place of the knocked out genes. We took our vial to the thermal cycler machine where it would perform PCR. This machine first heats up the sample to 98 degrees Celcius to denature the DNA, then cycles it through 25 to 35 cycles of temperature changes to perform annealing (adding the nucleotides), extension and a final hold to renature the DNA and finalize the process.
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At the end of Day 3, we left our vile in the thermal cycler. At the end of PCR, we will have a novel strand of DNA with no IRA2 and an antibiotic resistance gene in its place. This new strand will also have GFP fluorescence due to the use of the Nop1:GFP fusion strain that we used.
Day 4 - 9/7/16
Today we first learned about contaminants and how prevalent they can be in a variety of settings. In order to prevent such contamination as we started to create rich mediums for our yeast to grow on, we cleaned our lab area with ethanol and also lit 2 ethanol burners to sterlize the air. We then used the yeast given to us and micropipette it into four viles of YPD and two plates of agar, both rich mediums for growing yeast. We did this micropipetting close to the burners with brand new micropipette tips in order to keep the field as sterile as possible.
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In the second half of the lab, we performed gel electrophoresis on our PCR product. The reason we use gel electrophoresis is to look at the size of the DNA segments in order to see if the PCR was indeed successful. We can figure out how many base pairs long the DNA segment should be, then compare it to a ladder of known segment lengths through gel electrophoresis.
In order to perform gel electrophoresis, we took our PCR product and mixed 2 uL of it with 2 uL of a blue dye called GelRed (1:1 ratio) to make the PCR blue so that we could see it during the gel electrophoresis experiment. We then picked up the 4 uL solution and inserted it into a well in the gel electrophoresis machine. We actually did this twice because in the first well, we could not quite see the blue well enough. The second time we were more successful.
Day 5 - 9/12/16
Today we transformed our yeast through a protocol given to us by our professor. A link to this protocol can be found in resources. To begin, we once again cleaned our lab area and lit two ethanol burner to sterilize the area. The first few steps of the protocol were done for us. A colony of yeast was inoculated in 5 mL of YPD broth and grown overnight. The saturated culture was then used to re-inoculate 5 mL of fresh media and grown until it attained OD of 0.8. Yeast cells were then harvested by centrifuging at 6000 rpm for 5 minutes. This was what was given to us at the beginning of today's lab. We then removed the supernatant, or the solution in the vile by micropipetting it out. We washed the pellet that remained in the vile with 1 mL of water. This vile was then vortexed and centrifuged for 5 minutes at 6000 rpm.
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After this, we again removed the solution (which was water) through micropipetting. The protocol told us to use 200 uL of 100 mM of LiAc solution. We were given 1 M LiAc solution. In order to reduce that to 0.1 M, we had to mix 1 part (20 uL) of the 1 M LiAc solution and 9 parts (180 uL) of water. We then micropipetted all of that solution into the vile with our yeast pellet. This solution was then incubated at 30 degrees Celcius for 10 minutes.
After incubation, we vortexed the vile and mixed it in the centrifuge at 6000 rpm for 30 seconds. We once again removed all supernatant with the micropipette, leaving the pellet.
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We then added five solutions to the vile to resuspend the pellet. We first added 36 uL of the 1 M LiAc solution we used previously. We then added 25 uL of salmon sperm DNA. This helps the plasmid DNA that we will eventually add to get into the yeast. We then were instructed to add 50 uL of plasmid DNA and water. We originally had 50 uL of our plasmid DNA from PCR, however, we used 4 uL of it for the gel electrophoresis. Because of this, we added 4 uL of water to our plasmid DNA so that we would have 50 uL of solution total. We then added this to the vile with the yeast, LiAc solution and salmon sperm DNA. We also added 240 uL of 50% PEG and 30 uL of DMSO after that. We mixed all of the solutions and the yeast first with the micropipette and second with the vortex machine.
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The vile was then incubated again, first at 30 degrees Celcius for 30 minutes and then given a second heat shock by incubating it at 42 degrees Celcius for 20 minutes.
Day 8 - 9/21/16
Today we began working on developing a solid growth plate medium as well as figuring out the proper concentrations of our liquid YPD components. We spent the first half of the class working on an assigned worksheet about YPD and its components (yeast extract, peptone and dextrose) and optical density graphs. In the second half of the class, we made our YPD medium. We figured out, through some calculations, that we needed 2 g yeast extract, 2 g peptone and 4 g dextrose and water for 200 mL of YPD and the same plus 4 g agar for solid YPD (for our agar plates). We measured the ingredients using a mass scale and combined them in a large jar. Since we were concerned about contamination during this process, we then autoclaved our YPD mixture in its jar for 20 minutes.
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After the YPD was autoclaved, we poured 8 agar plates to be used late in our experiment. We also used some liquid YPD that was made by another group and poured 50 uL into four vials.
Day 10 - 9/28/16
Today we first got our plates back with the yeast that was grown in the 30 and 37 degree incubators for 48 hours. Unforunately, we discovered that the yeast must have been moved around a lot on the agar before drying because the colonies grew in very dispersed and random colonies that would be way too difficult to measure effectively (Pictures 11 and 12). Because of this, we found an empty area on the same plates and re-pipetted 10 uL of both yeast strains onto the plates. We were extra careful to pipette circular colonies and not move the plates before the yeast had dried. These colonies came out much better (Pictures 1 and 4).
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Next, we prepared the rest of our media for the experiments based on the groups we had established last class. For our caffeine and bleomycin groups, we had to heat up the agar we had prepared last class and mix it with the chemicals. We filled four 50 mL tubes with hot, liquid agar. Based on some calculations, we determined that in order to get 7.7 uM caffeine and 3.2 uM bleomycin, we need to add 75 mg of caffeine each to two of the 50 mL tubes and 12 uL of bleomycin each to the other two tubes. We then shook the tubes thoroughly to mix them up. Before the agar solidified, we poured these new medium into 8 plates, 4 caffeine and 4 bleomycin. We waited for these plates to solidify and then used the micropipette to put both yeast strains onto their respective plates. Because the agar had been mixed with the chemicals, the surface was not very smooth so we tried to put the yeast on an area that was relatively smooth to ensure the yeast would grow in colonies that were relatively easy to measure.
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After all of our plates had dried, they were once again taken to the incubators to grow.
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Today we also prepared our liquid YPD for the spectrophotometer. We made liquid YPD on Day 8, so we gathered this, both yeast strains and a 96 well plate to perform a serial dilution. For each yeast strain, we performed a 3 well serial dilution to dilute the yeast to 1/32 of its original concentration, as this is the concentration needed for the spectrophotometer. We did this by pipetting 64 uL of yeast into one well, then taking 4 uL of that and combining it in the second well with 60 uL of sterilized water to get to 1/16 of the original concentration. We then pipetted that solution up and down to mix it together. Then we took 32 uL of that solution (from the second well) and combined it in the third well with 32 uL of sterilized water to reach 1/32 of the original concentration. This same dilution was performed with both strains of yeast.
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Once we had the diluted yeast for both strains, we pipette 8 uL of yeast into 8 wells on another 96 well plate and then added 120 uL of the liquid YPD to each of those wells. This gave us 8 wells with 8 uL wild type yeast and 120 uL YPD and 8 wells with 8 uL mutant yeast and 120 uL YPD. This plate was then run through the spectrophotometer to obtain growth curves for both strains of yeast.
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Day 11 - 10/3/16
Day 12 - 10/5/16
Day 13 - 10/10/16
Today we had the intent of taking the second set of pictures of the colonies on our agar plates, however one of our TAs discovered that for some reason our plates had been thrown away into the bio-hazard bin and had been compromised. This was a very unfortunate discovery as a large portion of our data was now impossible to use since we only had the first set of pictures and could not determine a growth rate from only that data. While it was unfortunate, mistakes happen and we simply had to go off of the data that we had, which was the liquid YPD growth curves. As mentioned in the previous entry, the difference between the growth rates for the mutant and wild type strains was not statistically significant so those results were fairly inconclusive as well. However, we still discovered that the removal of IRA2 may not be enough to cause complete inactivation of Ras proteins and rapid growth in yeast cells. This does teach us something new about IRA2 and similar genes and their pathways in yeast.
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For the majority of the lab period, we did our final presentations for the biology portion of this lab and presented our data, analysis, results and conclusions. This PowerPoint presentation can be found in resources.
Day 6 - 9/14/16
Day 7 - 9/19/16
Today we learned about serial dilution, a technique involved in the measurement of the optical density of yeast. We first collected the necessary materials including ethanol burners, a 96 well microtiter plate, a micropipette and sterile tips, and both of our yeast strains. We pipetted 120 uL of our yeast strain into the first well. We then took one part of that and nine parts of water to fill the next well up to 100 uL (10 uL yeast, 90 uL water). We then took 10 uL of that diluted yeast and added 90 uL water in the next well and continued in this manner for 8 wells total. This procedure explains why this process is called a serial dilution. We also performed two more serial dilutions in two more rows of the 96-well plate. These dilutions were similar, however the second one was with two parts yeast and 8 parts water and the third was half and half (5 parts yeast and 5 parts water). This explains why in the graph of our results below there are three different lines. We ran our 96 well plate in the spectrophotometer to obtain the optical density graphs for each well. These results can be seen in the graph below. These serial dilutions and the spectrophotometer run were just for practice.
Today we presented an update of our lab experiences to our class. In this presentation (found in resources), we presented an updated hypothesis. After further research on the IRA2 gene and its function in yeast, we know hypothesize that due to our genes negative regulation of RAS proteins in yeast, its removal will result in the accelerated growth of cells. This is because the Ras/cAMP pathway that is regulated by yeast helps limit growth if it is occurring too rapidly or uncontrollably. Therefore, if IRA2 is removed or mutated, cell growth will be uncontrolled and the yeast cells will divide rapidly. We also found in our research that Ras proteins can be oncogenic, meaning that lack of their regulation could cause tumor-like growths.
We also discovered that IRA2 mutants have been shown to have increased sensitivity to certain toxins. Based on this information, our new hypothesis was if IRA2 is removed through genetic engineering, the mutant will experience rapid growth and increased sensitivity to toxins and therefore may be used as a model to study cancer in humans, since a similar gene/protein complex exists in humans and yeast. Ras proteins in humans are negatively regulated by the neurofibromatosis type 1 (NF1) gene and the GAP protein neurofibromin. Ras inhibitors are being studied as cancer treatments, so our experiment in this lab could potentially have some real world applications.
Today we started lab with a lecture from our professor about certain things we needed to include in the draft of our lab report for the biology section of this lab. The most important topic we discussed was the way we genetically engineered the yeast to create our mutant strains. There had been some confusion about what we used as template DNA and how we used PCR and the mechanism of homologous repair of DNA double strand breaks. Our professor cleared this up and explained that the template DNA was a yeast knock out collection strain with many genes knocked out and an antibiotic resistance gene put in their place.
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During the second half of the lab, we collected and analyzes our data from both the agar plates and the liquid YPD growth curves. We recieved our liquid YPD data from the spectrophotometer and started to plot the optical density for each well against time. We did this for each of the 16 wells that we used. The graphs can be seen below.
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To analyze this data, we calculated the slope of each graph at the point where the slope was greatest. We then averaged these slopes for the wild type and yeast strains and compared the averages between the two. In order to determine the statistical significance, we performed a t-test and calculated a p value using the averages, standard deviation and error for each group of data. The p value turned out to be 0.8638, indicating that the difference between the average slope of the wild type growth curves and the average slope of the mutant growth curves was not statistically significant. This did not support our hypothesis that the mutant strain would experience more rapid growth than the wild type strain. Some explanations for this could be error at some point during our experiments or another gene in the yeast strain replacing the function of IRA2. This is a very probable explanation because IRA1, a paralog of IRA2 with very similar function and/or MSI1, another negative regulator of the Ras/cAMP pathway in yeast, were not deleted and took over the function of IRA2 in the mutant strain. This would explain why growth was relatively equal in the mutant and wild type strains.
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We also got our agar plates back and started trying to figure out how to measure the colony sizes for the first round of measurements. We decided to take pictures from the same height with the same camera of each of the colonies, split the colonies into small areas and measured the height and length of those small areas on powerpoint. We then found the average height and length for the wild type and mutant colonies for all experimental groups. The pictures of these colonies can be seen below. Next class we will take the same pictures of the colonies and then compare the sizes against time to obtain a growth rate.
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Today was the first day of the chemistry portion of this lab. We first got an introduction from our chemistry professor about analytic chemistry and what it entails. Specifically, we learned about the techniques of microfluidics, PDMS, 3D printing and wax printing to create platforms that can then be used for measurement of yeast growth, in our specific case.
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In the second of the lab period, we did research on this type of chemistry and the specific techniques that were introduced to us. We specifically read many scientific articles of a famous analytical chemist named George M. Whitesides. As a group, we decided to combine wax printing, 3D printing and PDMS to create a platform to force yeast to grow in more controlled shapes or ways so that we can more easily measure its growth. We were also interested in whether or not yeast has the ability to grow vertically with or without contact with nutrients or nutrient rich media and whether or not gravity has an impact on yeast's ability to grow. In order to test out these questions, we decided to make a 3D model with horizontal and vertical channels, then cover it with PDMS and melt away the model so that we can test whether yeast grows more efficiently in the vertical or lateral direction. Next class we would focus on making this 3D model on tinkercad (a website for designing 3D models) and also designing a wax print experiment.
Day 14 - 10/12/16
For the majority of today, Morgan from our university library's digital imaging lab came in and told us about the 3D printer that the library has and how we should design and upload our 3D models to be printed.
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After her presentation, we spent some time messing around on tinkercad and starting to create a rough model of the 3D model we would use for the print.
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We also spent some time developing ideas on how we could incorporate wax printing in our experimental design. We decided to print white squares of different sizes on a black background so that when it is printed on the wax paper and the wax is melted, the white squares will be hydrophilic regions and the black will be hydrophobic so that agar can be put on those white squares and not cross into the black portion of the paper, thus creating perfect squares of agar to grow yeast on. The squares side sizes, from smallest to largest, are 0.508cm, 0.762cm, 1.016cm, 1.27cm, 1.524cm, 1.778cm, and 2.032cm. We want to see whether or not the yeast will grow up vertically on itself even if there is only nutrient rich agar at the base. This technique will also allow us to measure the exact dimensions of yeast colonies in order to compare their growth. We created our paper on word and it can be seen to the right.
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Next class we will print our wax paper, melt the wax and put the agar onto the white squares.
Day 15 - 10/17/16
Today we started off the lab period with the first round of presentations in the chemistry portion. Our presentation can be found in resources. We discussed our plans for our experiments including our 3D model, wax paper and use of PDMS. We also talked about what we would use to melt away the 3D model after coverin git with PDMS. A common substance used for this purpose is acetone, however, the 3D printer that we have access to prints using a polymer called PolyLactic Acid filament. This polymer can not be dissolved completely by acetone. After some research we found a few substances that could be used to dissolve PolyA including Weld-on #4 and strong bases like sodium hydroxide.
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After the presentations, we decided to print our wax paper model that we had created on word. A picture of the paper can be found below. We also continued to work on our 3D model on tinkercad and research how we will measure yeast's vertical and horizontal growth, specifically how we will measure the height of the colonies on the smaller squares of our wax paper if the yeast grows up onto itself as we predict it will.
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Next class we will finish up our 3D model, melt our wax paper and place agar on our squares.
Day 16 - 10/19/16
Today we split up the tasks of finishing up the 3D model on Tinkercad and performing the first practice trial of our wax print paper experiment. We finished up our 3D design, complete with a hollowed out sphere with a vertical and horizontal channel and a small channel to insert the yeast into the model. The way we made the channels was with a thin cylinder and a loop around it, forming a donut shaped hole that the PDMS can be poured in to create a channel. For the sphere at the base of the model, it is hollowed out and then contains another small sphere connected to the two cylinders of the channels so as to create a small space between the two spheres where the PDMS will fill in. A top view of a channel can be seen to the right. Once the model is printed, we will fill the two channels and the sphere with PDMS, then melt away parts of our 3D model including the inner cylinders of the channels and the inner sphere. We will use the strong base sodium hydroxide to melt away the polyA 3D printed model. We will not melt away the small cylinder on the side of the sphere because we will use this to insert the yeast into our platform. When the model was finished, we sent it off to Morgan so that it could be printed in the library over the weekend.
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For the wax print experiment, we took the paper that we printed last class and melted the wax using a hot plate. We placed aluminum foil on top of the paper to hold in the heat and melt the wax as efficiently as possible. We could tell when the wax was melted because the black ink has bled through to the other side of the paper. After melting the wax, we cut the paper up into smaller sections that would fit into petri dishes. In order to ensure that the agar would not flow directly through the porous paper, we applied a layer of a hydrophobic substance, Vaseline, onto the back of the paper sections. We then places the paper sections into the petri dishes and used a micropipette to carefully put the agar only on the white hydrophilic squares of the paper. We close up the petri dishes to maintain a sterile environment for the agar and finished for the day. Next class, if all goes well, we will put our yeast onto the agar squares.
Day 17 - 10/24/16
Today when we came in we took a look at our first trial wax paper petri dishes and found that they had some contamination. We realized that although the petri dish was fairly sterile, the paper was probably contaminated at some point in the process and after further research, we also discovered that vaseline is a food source because of its long carbon chains, so mold and other contaminants can easily grow on it. It was a likely a combination of these two things that led to the contamination we saw. We repeated the entire process, however this time we used a UV light on the wax paper pieces to sterilize them, and applied nail polish, another hydrophobic substance, to the back of the paper sections instead of Vaseline. Because of high demand for the UV light and nail polish, this was the only things we did during class today.
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Next class we hope to get our 3D model back and start with the casting of the PDMS onto the model. We will also put agar onto the squares on our wax paper.
Day 9 - 9/26/16
Today we learned about the use and application of the Florescence Activated Cell Sorting (FACS) machine. This is a machine used for cell separation by funneling cells out one at a time through a small vibrating channel and running a laser scanner through each individual cell for categorization. The machine then uses computational analysis to identify positively, negatively, and neutrally charged cells and the charges of cells are determined by what florescence they have in conjunction with their charge prior to coming in contact with the scanning laser. The machine then projects a shadow of the cell onto a screen so that the shadow sizes can be compared. Basically, the machine uses a small tunnel, laser and fluorescents to separate cells by size. This method of data collection could be helpful in some group's experiments.
Regular conditions
Caffeine
Bleomycin
Day 18 - 10/26/16
Today we used a pipette to put the agar onto the white squares on our wax paper with the dried nail polish on the back in the petri dishes under sterile conditions. Our setup can be seen in the pictures below. We balanced the UV light on two tip cases and placed the petri dish with the wax paper underneath. We also had two ethanol burners and a cleaned lab area where we would put the agar onto the white squares within the petri dishes. Before adding the agar, we once again put the paper under UV light to ensure that it was sterile. We then taped our petri dishes shut and together so that the environment within the dishes would remain sterile.
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In the second half of class, we gave our second round of presentations. In our presentation we restated our original hypothesis with few modifications and further explained the complications with contamination in our wax paper experiment. We also presented our ideas for measuring yeast's vertical growth and some applications of this experiment in the real world (or at least in the world of scientific research). This presentation can be found in resources.
Day 19 - 10/31/16
Today we worked on Tinkercad to make some adjustments on our 3D model because we found that when it was scaled down by Morgan so that the printer could print it, the dimensions got messed up and it did not print correctly. We also had to rethink our design because we realized that our hollowed out sphere would not work out given the way the 3D printer prints in layers. We made some adjustments including getting rid of the outer tubes and simplifying our model to a sphere with two cylinders. We shortened the two channels and made them thicker. We also made the small tube on the side bigger so that we can more easily pipette the yeast into our model. With this new simpler design, we plan to submerge our model into PDMS so that it forms a hollow ball with two channels and a third smaller channel to put the yeast in.
We also got our wax paper plates back and were happy to find that the nail polish and all of our other sterile techniques were successful in preventing contamination on the agar. Now that we had that technique down, we repeated the wax paper experimental procedure through the same process as last class, however, we had to figure out the correct amount of agar to put on each sized square in order to have the same proportion of agar:area on all the different sized squares. We performed some calculations to find the following volumes:
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We once again printed our wax paper, melted the wax on the hot plate, used the UV light to sterilize both sides of the paper and the petri dish and pipetted the appropriate amounts of agar agar onto the corresponding squares. One problem we faced was lots of bubbles forming as we pipette the agar so we had to scrape off the agar, UV light the paper again and try again. The technique that ended up being the most successful was pipetting with the tip almost parallel to the paper and only pushing the micropipette release button to the point of first resistance, rather than pushing it all the way down. We were mostly successful in pipetting agar onto the squares without bubbles.
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After we finished this process, we waited for the agar to dry, which did not take long, and then we figured out how much yeast to put on each of the agar squares. Since we repeated the wax paper experiment twice, we decided to put 20 uL on the first set of squares and 40 uL on the second set of squares. We pipetted these amounts onto the squares under sterile conditions and taped up the plates to be sent to the incubator.
Day 20 - 11/2/16
Today we got our wax paper plates back and examined the white squares for yeast. Unfortunately, we found no growth. We realized that we need to create an environment within the plate that prevents the agar from drying out. We researched some different techniques online and then decided the best and easiest way to keep our agar from drying out was to create a humidity chamber for the plates. The way we would do this is put the plates into plastic Ziplock bags with a wet cloth.
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Since we couldn't reuse the plates with the dried up agar, we repeated our entire wax paper experiment once again. We printed the wax paper, melted the wax on a hot plate, cut the paper up into sections that would fit into the plates, and painted nail polish onto the back of the white squares. Unfortunately, due to high demand for the UV lights and nail polish in the lab, this was all that we were able to accomplish today. We spent the rest of the day working on our next presentation and updating our website.
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Next class we will put the agar onto the paper once again, wait for it to dry, add the yeast to the squares, create the humidity chambers and see if yeast grows on the paper!
Day 21 - 11/7/16
Day 23 - 11/14/16
Day 22 - 11/9/16
Day 24 - 11/16/16
Today we picked up where we left off last class and started by applying UV light to our paper with the nail polish on the back. We then taped them down into petri dishes with tape that we alos put under the UV light. Then, using the micropipette, we put agar onto the squares in the same ratios that we used previously. We waited for the agar to dry (about 15 minutes) and added 20 uL of yeast to one of the paper trials and 40 uL to the second. We then created our humidity chambers for the petri dishes using large ziploc bags that we sterilized with ethanol and damp paper towels to keep the environment moist. We placed the petri dishes into the bags and sent them to be incubated at 30 degrees C for 48 hours. We are very hopeful that we will see yeast growth on this trial.
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We went to the library to check on our 3D print but it wasn't quite done yet.
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We spent the rest of the lab period finalizing our presentation for next class and researching the protocol for PDMS casting, which we will perform next lab period.
In the first half of the lab period today, we presented our group's recent work including our changes to our wax paper protocol and our updated 3D model design. This presentation can be found in resources.
In the second half of the lab, we got our plates back and were pleased to find that yeast had grown on quite a few of our agar squares. We created a sterile environment using ethanol burners and opened the petri dishes to take pictures of our yeast growth. We made sure to have the camera at the same height for the pictures so that they could be used for further analysis.
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Based on the growth we observed, we made the preliminary observation that the fifth largest square (2.322 cm^2 square) was optimal for the yeast growth. We made this observation because the growth on the square was closest to the edges of the square, even into the sharper corners, whereas the other growths didn't grow all the way to the edges or did not occur at all. It also appeared that little to no growth had occurred on the smaller squares so we hypothesized that the yeast did not have sufficient nutrients on those squares in order to have successful growth. In order to obtain a growth rate, we closed up the petri dishes, created new humidity chambers and sent the dishes to the incubator once again.
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Today we also performed our PDMS casting. We first picked up our finished model from the library and carefully ripped off the supports. We then made 22g of PDMS using 20g of PDMS and 2g of curing agent in order to create a 10:1 ratio. This ratio is optimal for our experiment because it makes the PDMS flexible by adding more monomers to create a long backbone so that it is easy to remove from petri dishes and our 3D model. We mixed the PDMS in a dish for about 10 minutes, then put our 3D model in a small petri dish and poured the liquid PDMS over it. Unfortunately, there was not sufficient PDMS left in our lab for us to completely cover our model as we had planned. We had to alter our experimental design a bit and decided to cover as much of our model as we could to create a sort of "cup" with the two channels protruding from it. We left the PDMS to dry overnight.
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Today we got our small petri dish with our PDMS and 3D model back and used a small metal probe to peel the PDMS out of the dish. There was some excess PDMS that we cut off with scissors. We then suspended the model in 1 M NaOH in order to melt away the polylactic acid filament of the model.
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While we let that sit, we took a look at our wax paper petri dishes that had been incubated for the second time. Unfortunately, we found that there was some contamination. We think this occurred because we used the same plastic ziplock bags as the first trial for our humidity chambers. Because of this, we were unable to use those dishes as data to obtain a growth rate for the yeast on the wax paper. However, we still had the pictures from the first trial and decided to repeat our wax print experiment in full, except this time only with the optimal square (2.322 cm^2 square). We printed out two sheets with only that size square, flowed the wax with the hot plate, applied nail polish to the back of the paper, applied UV light, then micropipette the agar and yeast onto the squares under sterile conditions. One one sheet, we put 20 uL of yeast and on the other we put 40 uL to optimize the amount of yeast used as well. We created a new humidity chamber (with new plastic bags and paper towels) and sent those petri dishes to the incubator.
Today we got our optimized trial of our wax paper experiment back and were pleased to find yeast growth once again. The yeast was not as easily visible in this trial for whatever reason, but the smell was undeniable. It smelled exactly like thriving yeast -- almost like beer. Based on what we observed, we concluded that 20 uL of yeast was much more optimal for the squares than 40 uL because on the 40 uL squares, the yeast was falling off the edges of the agar because there was simply too much solution.
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We also checked on our 3D print in the NaOH solution and found that it had barely dissolved at all. Unfortunately due to lack of time to try to dissolve the model a different way then add the agar and the yeast and observe growth, we had to end our 3D print/PDMS experiment at this step.
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We spent the remainder of class finishing up our final presentation, poster, and website.
Day 25 - 11/28/16
Today we gave our final presentations including a synopsis of our entire experiment and our conclusions based on the data we were able to obtain. This presentation can be found in resources. These presentations took up most of our class time and we spent the rest of the day putting some final details on our poster. The powerpoint version of our poster can also be found in resources.
Day 26 - 11/30/16
Today we had our lab practical in which we were tested our various laboratory skills and information that we have learned over the semester.
Day 27 - 12/5/16
Today we presented our posters to our professors and classmates. This was our final lab period of the semester and therefore this is the final entry in our virtual lab notebook! As mentioned previously, all final presentations and our poster can be found in resources.
