A few white colonies indicating the E. Coli cells kook up the hybrid plasmids were observed on the plate but no glowing colonies were detected. The lax Oberon was not successfully cloned in this experiment. Introduction Brio Fischer possesses a system called quorum sensing as a mean to express fluorescence and communicate collectively with other bacteria. (Chancier, 2011). Quorum sensing involves using signaling molecules called conductress transcribed from Lax I of lax Oberon for bacteria to coordinate their behavior depending on the environment (Enamel, 2004; Kaplan, 1985).
If the bacterial population density increases in the environment, semiconductors will accumulate and combines with the gene products of lax Oberon to produce fluorescence. (Rutherford, 2012) Lax Oberon contains seven Lax genes in specific orders. Lax CEASED are located downstream of the promoter and Luxury, which is transcribed in the opposite direction as all other genes, is located upstream of the promoter (Swartz, 1990).
When lax I- transcribed conductress accumulate and achieve a critical concentration in the environment, they will bind to the product of lax R and initiate the transcription and translation and produce luminescence effects. (Incongruent, 1983). Lax A and lax B are genes in the lax Oberon to transcribe the enzyme Lucifer that is essential to catalyst the fluorescence reaction with fatty acid reeducates complex transcribed from lax C, D and E. (Neighed, 1988). The overall purpose of this experiment is to create a genomic library and clone the LULAS genes from the marine bacterium Brio Fischer by using Recombinant DNA.
Restriction enzymes are used to digest the genomic DNA and create DNA fragments as inserts. The inserts are then combined with the vectors and legated by aliases in order to transform the recombinant 2 DNA into the host. The ultimate goal is to express the lax Oberon gene products to produce fluorescence in another organism, E. Coli that is not capable of expressing the light naturally. The article “High-efficiency transformation of mammalian cells by plasmid DNA” uses a similar approach to clone a specific gene. Chin, 1 987) We hypothesize that if the hybrid plasmids containing lax Oberon is successfully transformed into the E. Coli DUH-alpha cells, the cells will produce fluorescence and glow in the dark. The biggest implications that will impact the success of this experiment is the possibility that we will lose the gene f interest during the process. Therefore it is important to use gel electrophoresis to check if digestion and legation work to ensure the gene of interest is being cut and inserted properly.
Material and Method Part I: Isolation of genomic DNA of brio Fischer. The culture of V. Fischer is first centrifuged to separate the cells from supernatant. The following reagents are added to destabilize the outer membrane of the cells and denature the proteins: TEST buffer, loosely, proteins K and SD. Phenol is then introduced to separate the proteins and lipids from the nucleic acids. Ethanol is added after removing the aqueous phase of DNA to aggregate DNA structures. Use a hook to wind the DNA and turn it upside down to drain and get rid of all the ethanol.
Place the DNA into ET buffer again and add Rinse and proteins K to discard RNA and further purify DNA. Phenol and chloroform are introduced to further extract chromosomal DNA. Centrifuge and pipette the aqueous layer out and add sodium acetate and ethanol to further aggregate the DNA Use the hook to wind the DNA out from the solution and dry it. After several minutes, dissolve DNA in ET buffer. To check the purity of DNA, the spectrophotometer is used to measure the absorbency readings of the chromosomal DNA sample at both wavelength of 260 NM and 280 NM.
Use the ratio of 260 to 280 to calculate the DNA purity and determine possible contamination. Part II: Restriction digest of the genomic DNA and page vector with Sal. 3 The genomic library is created by using restriction enzyme Sal I to digest the DNA into fragments of different sizes. These fragments will later be inserted to the plasmids and transformed to E. Coli DUH-alpha cells. Four samples are prepared or gel electrophoresis to test if the digestion works: Undigested brio DNA with Sal l, undigested brio DNA alone, undigested page plasmid DNA with Sal I, and undigested page plasmid alone.
The Dana without Sal I are served as negative controls to see if digestion works. The lanes loaded with DNA and Sal I should show several bands that represent linear conformation of the DNA. For page plasmids in particular, the linear band will have approximately 3200 base pairs. The lanes loaded without Sal I will have either band representing circular conformation (for brio chromosomal DNA) that stays very close to the well or overcooled conformation (for page ) that travels furthest away from the well.
Part Ill: Legation of the genomic DNA into page vector. Enzyme aliases is used to legate the insert and the vector. The insert to vector ratios are 1: 1, 2:1, 3:1 and 4:1. To test if legation works, load solution containing different ratio of inserts and vectors into multiple lanes with addition of aliases (lane 2, 4, 6), and solution containing inserts and vectors without aliases next (lane 1 , 3, 5). Then load one lane with digested plasmid and aliases (lane 7) and one lane with digested plasmid only (lane 8).
Digested plasmid without aliases serves as a negative control for lane 1, 3, 5 to observe circular bands or supercilious bands on the gel; Digested plasmid with aliases serves as a negative control for lane 2, 4, 6 to show linear bands that indicates successful legation. Part ‘V: Transformation of the legation products into competent Escherichia Coli Cells. E. Coli strain DUH-alpha cells are first made competent for them to take up recombinant DNA. The competent E. Coli cells are then added to the legation products. Heat shock the solution for a short period of time, then add the nutrient LB.
Divide each samples into different volumes and Spread the transformed cells onto plates containing LB, inclining and Gal. Use 4 blue- white screening to identify colonies that take up the recombinant DNA and examine the plates in a dark room for glowing colonies. Negative control is done by adding ET buffer on to the plate to ensure no colony growth after incubation. Spread the solution containing uncut page plasmids with competent sells on another plate to calculate transformation efficiency. Results DNA purity and concentration The DNA absorbency at wavelength of 260 NM and Mann is 0. 77 and is 0. 79, respectively. We calculated the extracted DNA purity by using the ratio of , which is = 1. 79. Brio chromosomal DNA Concentration was calculated by first multiplying AWAY of the sample with dilution factor: 0. 677 x 10 = 6. 77 then using the following equation: 1 AWAY = 50 to find the DNA concentration (x): ? : x = 6. 77 * 50=338. 5 . 0. 3385 = 0. 339 . Restriction digest of the genomic DNA and page vector with Sal. Due to absence of the bands from lane 1 and 2 in figure 1, digested DNA concentration could not be estimated based on the gel results.
Thus the concentration is calculated y using the amount of DNA from the solution containing Brio DNA and Sal I divided by the total volume of the solution. 1 Quo, or 10,000 Eng of Brio DNA is loaded in the solution with total volume of soul. The digested DNA concentration 5 = 200 In figure 1, Lane 3 shows multiple bands which represent page DNA plasmid in different conformations. The first band that is the closest to the well is linear form of the plasmid, which is approximately 3200 base pairs and represents digested plasmids. The smeared band represents supercilious conformation representing uncut page.
Digested page DNA concentration can be estimated y comparing the intensity of linear bands in lane 3 with standard Mind Ill bands. The intensity of the linear band is similar to the third band of the standard, which has 6557 base pairs. Thus the amount of DNA in the linear band can be calculated as the following: x 500 Eng = Eng of DNA. Since lull of PAGE was loaded into the well, the concentration of digested page is = 35 6 Legation of the genomic DNA into page vector. In figure 2, all the lanes show circular bands pattern near the 23130 BP standard Hind Ill bands indicating genomic DNA that are not being cut.
In lane 1 and 3, which are loaded with existed brio DNA and plasmid page DNA without any aliases, show faint bands in between 4361 base pairs and 2322 base pairs in accordance with the standard ladder representing linear conformation of page plasmid; Lane 2 and 4, which are loaded with the same Dana as lane 1 and 3 but with addition of the aliases, show more clear view of supercilious bands with the same level as band 5 of the standard ladder. Due to absence of negative control lanes that are loaded with plasmids only and plasmid with aliases, it is difficult to determine if legation is successful. Transformation of the legation products into competent Escherichia Coli Cells After incubation, numbers of blue and white colonies on each plates are shown in table 1. Iii and lull of recombinant plasmids from sample 1 with insert to vector ratio of 1. 45: 1 have white colonies as well as lull of recombinant plasmids from sample 2 with ratio of 1. 66: 1 . No glowing colonies are observed. Plate TO served as a negative control of the transformation experiment showed no colonies on the plate. The white colony percentage indicates which insert to vector ratio yields better transformation.
The transformation efficiency can be calculated by using TO solution, which contains lull of uncut page plasmid with incineration of 0. Eng/LU, lull competent cells and iii of nutrient LB to figure out the number of colonies per one micrograms of DNA. The amount of page is lull (0. 2 Eng/LU) 1 Eng. As we took Lou from the above TO solution (1 Lou), we could calculate the amount of page within that lull that we took out by the following equation: = X Eng of page = Transformation efficiency = GU colonies/ lug plasmid DNA. =O. Eng=xx When we took out 1 iii from the TO solution, we also calculated the transformation efficiency with the same method: = X Eng of page Transformation efficiency = DNA. = O. Ins = 1 x GU colonies/ lug plasmid 6. 1 x 8 Transformation TTL page 950 LB 45 LU competent Cell TO page 950 LB 45 LU competent Cell volume Plated WI) 10 50 200 10 50 200 Total Colonies 129 761 84340247 520 # Blue Colonies 129 752 840 40 247 512 # White colonies 0 93 0 06 % white colonies 1. 15% Glowing colonies 00 00 0 0 TO lull sterile ET 950 LB 200 00 0 00 45 LU competent cell TO 10 8 8 0 0 0 lull uncut page BIBB lull competent 100 61 61 00 0 cell Table 1: E. Oil colony counts. The table shows the numbers of blue and white colony counts as the results of transformation of competent E. Coli DUH- alpha cells with recombinant plasmids. The plates are incubated for 18-20 hours at 37 Celsius. 9 Discussion DNA extraction DNA purity can be determined by the ratio of Mann over 280 NM since DNA maximally absorbs the light at wavelength of Mann and proteins absorb the light at wavelength of Mann. The ratio for a pure DNA solution is 1. 8. If there is protein contamination in the solution, the ratio will drop to below 1. ; On the other hand, if there is a lot of RNA contamination in the solution, the ratio will be above 1. 8. The calculated DNA purity of the sample in the experiment is approximately 1. 79, which is close to the 1. 8 standard and indicates good DNA purity. However, since the ratio is still below 1. 8, we do not rule out the possibility of protein contamination. There are several methods we can adjust during the process of DNA extraction to decrease the chance of protein contamination.
First is to be more careful when petting out the aqueous layer from the proteins and all other organic molecules; Secondly, we can add more chloroform to make the interface layer in between the aqueous solution and organic layer clearer. It is also helpful to centrifuge more frequently to ensure each layers are separated. Restriction digest of the genomic DNA and GEM vector with Sal. The first gel electrophoresis is run to ensure that brio DNA and the plasmids are being digested by the restriction enzyme Sal l.
If brio DNA is being digested successfully, we expect to see multiple bands across the lane since Sal I may create multiple genomic DNA fragments with different base pairs; In comparison, lane 2 is expected to have a band that appears very close to the well with the same level as the first band of the ladder, indicating genomic DNA that is not being cut and remain its circular conformation. If the plasmids digestion is successful, we expect to see a band close to the size of 000 base pairs representing linear conformation of the plasmid in the lane that contains Sal l.
The negative control lane which has no Sal (lane 4) should have supercilious band that travels further away from the well than the linear band, indicating none of the page plasmid is being cut and the digestion occurred solely due to the addition of Sal l. However, in the lane 4 of figure 1, which we expected to see supercilious bands only, also shows a band with the same distance as the linear band in lane 3. This indicates possible cross contamination when we are loading the gel, and lane 4 fails to serve as a negative 10 intro to indicate digestion success.
To improve gel reliability, we should remake the solution to ensure that no Sal I is added with the page plasmid in the negative control solution and re-run the gel again for better interpretation. In addition, the picture of the gel is being cut off since the first band of the standard ladder is 9416 base pairs instead of 23130 base pairs. The reason why the first band is determined to be the 9416 base pairs is because in a normal picture of gel with Hind Ill as the ladder, the 4th band (4361 base pairs) always has the least intensity compared to the first 3 bands and the 5th bands.
In figure 1, it is actually the third band being the faintest, which leads to the conclusion that the third band is in fact 4361 base pairs and thus the first band is 9416 base pairs. Without an intact ladder, the interpretation of bands and their intensities become less reliable. It is important to always check the picture first before throwing away the gel. The biggest reason why figure 1 fails to indicate the result of brio DNA digestion is due to the missing bands in both lane 1 and 2.
Without any bands, we could not predict whether our genomic DNA is being successfully existed with Sal I or not, and we could not conclude whether Sal I is the reason why DNA is being digested. There are many reasons to explain why no bands were shown in lane 1 and 2. First one is the picture being cut off. It is possible that the bands are very close to the well and on the same level as the first band of the ladder. However without having a complete picture of the gel, we can’t simply explain whether the DNA is successfully loaded or not.
The solution to this problem is, as addressed earlier, to retake the pictures; Second reason is due to small concentration of genomic DNA. According to results, the DNA that s extracted from the Brio Fischer cell has only 0. 339 GU/LU concentration. The solution is too diluted that increases the difficulty of transferring DNA form the solution to the gel. To solve this problem, we can either enhance the DNA extraction skill by petting more aqueous solution out to increase DNA concentration, or to vortex the DNA solution before petting into the gel so DNA is well distributed throughout the solution.
It is also possible that the genomic DNA has residual protein contamination that may possibly inhibit Sal l. This shows how important the extraction step is and how the very beginning steps an hugely impact the results of the experiment. We compare the intensity of the bands on the gel representing digested Dana with the ladder’s to determine the concentration of the digested Dana. In lane 3 of figure 1, we conclude 11 that the linear band, which is the band that’s on the level between 4361 BP and 2322 BP of the ladder, has approximately the same intensity as 4361 base pairs.
Thus the amount of digested page plasmids has approximately the same amount as the 4th band of [II-Hind Ill, which gives us an indication to calculate the actual concentration of the digested DNA. The calculation is shown in the exult section. Since the genomic DNA concentration is so low and the digestion gel fails to indicate if DNA is cut or not, we decided to put in as much volume of our digested DNA solutions as possible to increase the possibility of making the recombinant DNA. We did this to ensure that all the digested DNA we have can be inserted into the plasmids.
The first sample contains 4. Jug digested inserts with 2. Jug digested plasmids, so the ratio is 1. 45: 1; The second sample contains 4. Jug digested inserts with 2. Jug digested plasmids, so the ratio is 1. 66: 1 . Legation of the genomic DNA into page vector. The second gel is run to check if the legation works. When aliases is introduced to the solution with digested inserts and plasmids, the enzyme will start legate every ends of the fragments that are cut by Sal I with sticky ends.
There are many combinations produced by aliases, such as inserts legated to inserts, inserts legated to the vector, inserts that is our gene of interest legated to the plasmids, or plasmids re-legated themselves. Figure 3 shows our prediction of legation gel. Without the addition of legation, digested inserts would not be legated with the digested plasmids, thus plasmids main in linear conformation and the lanes without aliases addition should have a band with size of approximately 3200 base pairs.
Lane 5 serves as a negative control to show the band pattern of a cut plasmid. By comparing lane 5 with lane 1 and 3, we anticipated that lane 1 and 3 both would have a band similar to lane 5 to indicate the lane is loaded with digested plasmids only; Lane 6 serves as a positive control to show the supercilious conformation the plasmids become after the addition of aliases. We expect lane 2 and 4 would have the same supercilious bands as lane 6 to show that the legation does work and the inserts are successfully legated with the plasmids.
Lane 6 also would have supercilious bands that are further away from the well than the supercilious bands from lane 2 and 4 due to the fact that the plasmids re-legated itself in lane 6 (the size is approximately 2000 base pairs) while the plasmids take up the inserts in lane 2 and 4 so the size of the plasmids will be bigger. Lane 1 and 3 also serve as 12 controls to for lane 2 and 4 to show the difference between digested plasmids versus plasmids that are being legated. There are also other patterns of bands that would likely be shown on the gel that are not shown from figure 3.
In lane and 4, it’s possible to also see linear band which indicate that aliases does not legate all the plasmids in the solution; It’s also possible to see a supercilious band that’s at the same level as lane 6, which indicates that there are some plasmids that are being self-legated in both lane 2 and 4. Lane 1 and 3 will possibly have multiple linear bands due to different sizes of digested genomic DNA and the plasmids. However we expected Lane 1 and 3 to have at least one linear band that’s the same level as the lane 5 to ensure the digested plasmids are being loaded to the lane correctly.
Figure 3: Anticipated gel results of legation. Lane 1: digested brio DNA + digested plasmids (1. 45: 1) Lane 2: digested brio DNA + digested plasmids (1. 45: 1) + aliases Lane 6: digested plasmids + aliases Lane 3: digested brio DNA + digested plasmids (1. 66 : 1) Lane 4: digested brio DNA + digested plasmids (1. 66: 1) + aliases Lane 5: digested plasmids 13 However, the actual legation gel (figure 2) fails to show that legation works due to several reasons. First, in figure 2, no bands were shown in digested plasmids (lane 5) and digested plasmids with aliases (lane 6).
Without having these two as controls, we could not conclude the band patterns in lane 1 to 4. The reasons eight be because the samples of lane 5 and 6 were sunk to the bottom of the gel so they were never traveled across the gel during electrophoresis; In addition, it is also possible that the background of the picture gel is too that the bands become difficult to visualize. The solutions to these two possible reasons are to add less Sybil safe into the agrees gel so the background can be less green; Also, if possible, reload the samples and run the gel again to have a better picture before moving on to the next step of the experiment.
In addition to the controls, it is also very difficult to visualize the expected bands in lane 1, 2, 3 and in figure 2. Lane 3 and 4 have really faint bands that are most unlikely to see without staring at the picture for a longer period of time. Lane 4 has a possible supercilious band at the level of 2027 base pairs, while lane g’s expected linear band is almost non-existent. Lane Xi’s expected linear band is better to see and the supercilious band in lane 2 is also visible.
Other than the possibility that the background is too green that affects our ability to see the bands, it is also impossible that the amount of Dana in each well is too low for the band to be visible. As addressed earlier, the DNA we extracted has low concentration and his may decrease the amount of digested DNA in the sample solutions. To solve this problem, we should improve our DNA extraction skills to increase the overall genomic DNA concentration. Besides the DNA concentration, there are also other possibilities that may affect legation of inserts and the plasmids.
One is that only a small amount of DNA is being cut by Sal I in the earlier step of the experiment, thus only a small amount of inserts and vectors are being legated by the enzyme; Secondly, it is possible that most plasmids are being re-legated back so less amounts of inserts are actually being taken up. One way to solve this problem is to use alkaline phosphate, which takes away the phosphate group from the page causing page not able to re-legate. This increases the likelihood that the inserts can be legated with the vectors.
The third possible reason is failure to inactivate restriction enzyme that were previously added into the solution. Without inactivating Sal l, it is possible that the plasmids and the inserts are being cut again after legation. 14 In conclusion, the gel did not sufficiently provide us an answer regarding legation. This step should be repeated again before moving on to transformation. Last part of the experiment is to apply blue-white screening method to detect glowing colonies. We are expected to see blue colonies and white colonies on the plate.
The page plasmid is engineered to include a Lace-Alpha gene in the multimillion site. When the plasmid is taken up by the E. COli cells without any insert, the plasmid will transcribe the Lace-alpha subunit. This alpha subunit will combine with the omega subunit produced by the E. Coli cells and activate beta glaciological protein to breakdown Gal and make the colony blue. However, if the genomic DNA is inserted and legated with the plasmid, the Lace-APIPA gene ill be disrupted and thus no alpha subunit will be produced inside the cells.
Without the alpha subunit, beta-glaciological will not be activated and the colony will remain white. This method helps us differentiate colonies that took up a plasmid with an insert, which shows white colors, or plasmids that did not take up any inserts, which shows blue colors. If any bacterial cells took up the gene of interest, the lax Oberon, the colonies will glow in the dark. According to table 1, we are expecting TTL plates and TO plates to have blue and white colonies and maybe even glowing colonies.
TO is served as a negative control to ensure hat colonies grown on the media are solely due to competent cells taking up the plasmids; TO plate is served as an indicator of transformation efficiency. We loaded different volume of the solutions onto TTL and T 2 plates and calculated white colonies percentage in order to find which insert to vector ratio yields the most white colonies. Table 1 shows that TTL plate, with insert to vector ratio as 1. 45:1 and Iii plated volume, yields the highest white colonies percentage (1. 18%).
However, this is not a good reference for future experiment because the insert to vector ratios between two samples are really close to one another. The experiment should be repeated one more time to ensure the consistency. Transformation of the legation products into competent Escherichia Coli Cells We did not observe any glowing colonies at the end of experiment, which indicates that the gene of interest is lost during the experiment and is not taken up by the plasmids and transformed into E. Coli cells. The gene of interest could be lost from the DNA extraction step, since the final DNA concentration we extracted was only 0. 39 GU/LU and the gene of interest may be left with the aqueous solution that we discarded; It is also possible that the gene of 15 interest was never taken up by the plasmids during the legation process, thus never being transformed into the bacteria. The transformation efficiency calculated in the result section indicates the skill of transforming the competent cells with plasmids onto the media. According to the article “High efficiency transformation of Escherichia Coli with Plasmids” (Inure, 1990), a good transformation efficiency is between to colonies/GU.
A lower transformation efficiency values indicates that the technique should be improved. The calculated transformation efficiencies from TO with two different volumes, lull and 1 iii, according to table 1, sis x colonies/ lug and 6. 1 x colonies/ lug plasmid DNA. We expected these two values to be close to each other to ensure technique consistency; however, the efficiency is considered lower and one possible reason is the competent cells being damaged during the process of transformation.
We need to make sure to pipette the competent cell solution very carefully without any bubbles to increase our transformation efficiency. Conclusion We hypothesize that if the hybrid plasmids containing lax Oberon is successfully transformed into the E. Coli DUH-APIPA cells, the cells will produce fluorescence and glow in the dark. However, we did not observe any glowing colony at the end, thus we were not able to amplify the gene that produces fluorescence and sequence to see if it is lax Oberon.
One way we can improve this experiment to increase the probability of having glowing colonies is to load the digested brio DNA samples into the gel and separate all the inserts into three different group of fragments ( one group that has 6500 base pairs of more, second group that has 3 Kbps to skip, and the third group that has the lowest sizes, etc). Then take the Dana out from the gel and perform legation separately. This method can increase the likelihood of inserting the gene of interest with the plasmids.
The discovery of lax Oberon brings tremendous advantage to research in molecular biology. The fluorescent light can be used as a genomic marker to tract the gene or protein of interest or to monitor gene expression in different organisms. By using the correct restriction enzymes and aliases and carefully examine every steps of the experiment, the lax Oberon will be successfully transformed into the target cells and produce fluorescence. 16 Reference 1. Aimer, B. M. M. 2004. Cell-to-cell signaling in Escherichia coli and