Gibson Assembly Attempt with 7kb and cmr

 Introduction 

Previously, PCR amplification was run on 7kb plasmid extracted via the Zyppy Kit and cleaned up through the Monarch PCR & DNA Cleanup Kit (5μg). When viewed on a GelRED agarose gel, 3 bands were present, 2 were non-specific to the 7kb we wanted. However, we will continue to attempt Gibson assembly using 3 times as much vector to make up for 1/3 of our sample being the 7kb plasmid. Afterward, we will transform competent E.coli cells with the Gibson assembly product. 


Methods 

To attempt Gibson assembly, we began by diluting our 46ng/μl cleanup sample of cmr to a concentration of 11.5ng/μl. We diluted our sample to keep our vector and gene equimolar. Next, on ice , we added 9μl of our 7kb samples to 1 PCR tube each. 1μl of our diluted cmr sample was then added to both tubes, making a total volume of 10μl in each. Next, 10μl of Gibson assembly mastermix was added to each tube. Our positive control had 10μl of control reagent and 10μl of Gibson assembly mastermix. All tubes were  placed in the Thermocycler and incubated at 50°C for 15 minutes. Lastly, samples were stored at -20°C.


To attempt transformation, competent E.coli cells were thawed on ice. We then chilled 2μl of ligation mixtures in 2 1.5mL centrifuge tubes. Next, we added 50μl of competent E.coli cells to the DNA and mixed via resuspension and flicking. These mixtures were then placed on ice for 30 minutes. To heat shock, cells were placed in a 42°C water bath for 30 seconds then placed on ice for 2 minutes. Immediately, 950μl of LB media was added to each tube. We then placed these tubes in a shaking incubator at 37°C for 60 minutes. Lastly, 50-100μl of our mixtures were spread on pre-warmed selective plates. Our 4 variant selective plates had LB with no antibiotic , chloramphenicol at 25μg/mL , chloramphenicol at 170μg/mL, and ampicillin at 50μg/mL. Our 4 variants were our positive control mixture, our negative control containing no plasmid, and our two experiments containing different extraction samples. There were a total of 15 plates streaked with 50μl



Results 

Gibson Assembly Plasmid #1 , no growth on chloramphenicol plates



Gibson Assembly Plasmid #2, no growth on chloramphenicol plates


Postive control, growth on Ampicillin and NA plate


Negative Control, no growth on antibiotic plates, growth on NA plate



Discussion

No growth of our Gibson Assembly transformations were seen on chloramphenicol plates. Therefore , we can conclude something went wrong during the Gibson assembly reactions as E.coli cells are extremely competent. Our next steps would be to find a more suitable PCR reaction. Our 3 bands after PCR cleanup likely are the reason we weren't able to have successful Gibson assembly. Finding better PCR settings would ideally require a heat gradient mixed with variable MgCl additions. However, due to lack of plasmid extraction template, four reactions will be ran. 





Comments

Post a Comment

Popular posts from this blog

Colony PCR Amplification of 7kb Plasmid from Deinococcus aquaticus and cmr from E. coli's pRAD1 Plasmid

Image
 Introduction  This semester, Leilani Boren, Micheal Dockham and I are transforming the 7kb plasmid from Deinococcus aquaticus with E.coli's chloramphenicol resistance gene via PCR amplification and Gibson assembly. Once assembled, the 7kb plasmid will be farmed and reintroduced into D. aquaticus. To do this, we must first prime D.aquaticus' 7kb plasmid as our vector and E. coli's chloramphenicol resistance gene as our antibiotic resistance gene via PCR.  DNA cloning works by inserting a target gene (cmr) into a plasmid (7kb) . Afterward, the plasmid is introduced into bacteria (E.coli) via transformation. Colonies will be screened via selective media (LB + chloramphenicol) and, once verified, will be used to farm recombinant plasmid DNA.  Typically, Plasmid DNA is manipulated via restriction enzymes which will cut at cloning sites to create an area for our fragment to be introduced and hybridized. However, we will be using the Gibson Assembly due to successes in combining
Read more

Restriction Enzyme on Deinococcus Aquaticus 7kb Fragment

Image
 Introduction Last week in lab, two largely concentrated samples of the 7kb fragment from Deinococcus Aquaticus was obtained. However, the gel showed this sample to be at 14kb. A theory for this is that two 7kb fragments are mineralized together. Therefore, we are using restriction enzyme to demineralize these fragments and have them appear at 7kb. If this isn't the case, we will use primers. Restriction enzymes are sequences that are recognized by restriction enzyme to cut plasmids. We cut at the restriction site xbaI (1188 base pairs) but this should show as 7kb on the gel.  Procedure A 1mL tube was filled with 68ul pcr water, 10ul cutsmart buffer, 2ul restriction enzyme , and 20ul of our plasmid isolation samples from the week prior (2mg). Once we have a 100ul total , we will incubate this tube on a heat block at 37 degrees celcius. Due to not knowing how long this enzyme will take to completely cut at all of the restriction sites, we will use 4 incubation time periods. These ti
Read more

Using New GelRed on Edited 7kb PCR Reactions and Extractions

Image
 Introduction  Recently, a heat gradient from 50 °C-66 °C   on the 7kb plasmid from D.aquaticus was ran and contained no visible banding. Due to an empty agarose gel, this may reveal that the template we're using ( boiled cells ) doesn't contain the 7kb plasmid. This would mean our primers aren't able to bind appropriately. Another possibility is that our annealing temperature. Therefore, the alternative bead-beating template will be used in a PCR reaction and run on the more sensitive GelRed agarose gel.  Additionally, a new lysing protocol involving proteinase k and lysozyme will be used to try to extract 7kb plasmid. A theory to test is that we've obtained 7kb plasmid and hadn't seen bands due to the previous dye needing >500ng of template DNA. This new GelRed should only need 5-50ng of DNA to bind visibly.  Methods To run LongAmp PCR products, PCR settings were set to the following: initial denaturing was set to 4 minutes at 94 °C, denaturing for 30 seconds a
Read more