Plasmids - Science topic

Hello all,

I have a mini-prepped DNA that I am trying to digest with EagI. After doing the digestion it looks like the reaction with the restriction enzyme maybe linearizing my plasmid as it migrates at its expected weight when next to the DNA ladder. However the undigested sample runs very high, far above the 10kb band of the ladder. I am guessing this is the uncut is showing a nicked plasmid, which is why it's running so high? What do you think?

Thank you

MB

I have designed and assembled a 10kb plasmid containing SbfI and NcoI restriction sites flanking a promoter region, which I am attempting to excise and replace with other promoters. When I do a double digest with both enzymes, I do not get my expected 750 bp band. I have already sequenced my plasmid and it definitely contains both of my restriction sites where I am expecting them.

I have digested my plasmid with SbfI and NcoI individually several times and it looks like each enzyme is working as I am seeing a linearised product on an agarose gel. Both enzymes are HF supplied by NEB and follow the same protocol with the same reaction conditions.

I've tried running the reactions for longer doing 30 min, 1 hr, 3 hr digests. I've used DNA from different (although Sbf is unstable after 1 hr) minipreps, different quantities of DNA (0.5 to 5 ug), different volumes of SbfI (0.5 to 3 ul), brand new restriction enzymes and rCutsmart buffers.

I've included a gel image below. Using a 1% agarose gel ran at 90V for 45 min and a Quick Load Plus DNA ladder.

I don't know what else to try or what else could possible be going wrong.

Any help would be greatly appreciated.

Hello

I am currently engaged in research focusing on plant genetic transformation. As part of this endeavor, I have designed a comprehensive in silico plasmid cloning approach. The aim is to enhance the accuracy and efficiency of our genetic engineering efforts in plants.

I welcome and deeply value input and suggestions from all stakeholders involved in this research. Your insights and expertise are crucial as we strive to optimize our methodologies and achieve impactful results in the field of plant genetic transformation.

Together, through collaboration and exchange of ideas, we can ensure the effectiveness and precision of our approaches, ultimately advancing our understanding and application of plant genetic engineering for various beneficial purposes.

Anyone is invited to write me at [email protected]

Thanks in advance everybody

A plasmid of mine has GOI in opposite orientation. I have another plasmid which has exact same RE site but in such a way that it orients my GOI in right direction. However even after multiple attempts of RE digestion, Gel extraction followed by ligation, I didn't got any colonies on my transformation plate. I also checked efficiency of my competent cell with empty vector and found no issue (got 200-300 colonies) . I would be grateful if anyone can suggest me any possible solution.

Hi,

I'm trying to find a pDNA transient transfection carrier for my MDA-MB-231.

I'm using either liposome or lipopolyplex, but the transfection variability between experiments are too great.

I can only suspect that thin film hydration method has high variability (due to water bath sonication) and changed to ultrasonication which gave 0% transfection (positive ctrl worked, so no probs with pDNA).

Here's the protocol.

1) Thin film made in 4-mL vial or RB or e-tube using rotovap (5 mg/mL lipids in chloroform)

2) Hydration tested with DW/opti-mem/PBS/HEPES using water (1 mg/mL)

3) DNA solution added to liposome solution while vortexing

4) Cells treated with 1 ug DNA/well in opti-mem for 4 hrs, then complete media exchanged or added

For lipoplexes,

1) LPEI solution was added to DNA solution (N/P=10), RT incubation, 30 min

2) Liposome mixture thin film made as above

3) Hydration tested with various buffers or polyplex solution

4) Polyplex solution added to liposome

5) Cells treated with 1 ug DNA/well in opti-mem for 4 hrs, then complete media exchanged or added

I'm already on a number of tries and been frustrated with the result because no matter how consistent I am, the results are different.

Please share your wisdom with me!

In particular using a DNA plasmid transient transfection on differentiated skeletal muscle cells?

I am trying to clone the cowpea gene (3.5Kb) under the control of the 35s promoter (Gateway plasmid pK7WG2D). I used pDONR222 for the BP reaction. I got the positive colonies, and then sequencing confirmed that my gene was inserted in the donor plasmid. After the LR reaction, I got a lot of positive colonies in the antibiotic (Spectinomycin) plates. But when i do the colony PCR or PCR after plasmid extraction, i don't get amplification with my gene specific primers. However, I get amplification while using backbone-specific primer. I tried multiple protocols and used top10 and DH5a for transformation, but I had the same issue. What could be the problem?

Can anyone share their experience regarding the acquisition or gifting of cell lines mentioned in a Nature article? Specifically, I am interested in knowing if there are any journal-specific policies involved.

The cell line in question contains a CRISPR-mediated stably expressing protein labeled with GFP, and its published in the Nature journal. I intend to use this cell line for my own experiment. Naturally, I am willing to acknowledge or provide authorship as appropriate. However, I would like to know if it is possible to obtain this cell line directly from the PIs lab and what the relevant policies are of nature journals if I get them from PI lab (I understand I will refer the article).

Has anyone ever received or gifted cell lines before? I am aware that exchanging plasmids is a common occurrence, but I have never personally obtained cell lines in this manner.

I appreciate any information you can provide.

Thank you in advance.

I would like to track in real time the growth of bacteria using fluorescent miscoscopy. I remember there was at least one plasmid that could be used: transfected bacteria would emit a red or green fluorescent signal that could be used to track their growth and position.

Alas, I don't remember what was the name of these plasmids and I can't find a reference in the literature.

Does somebody know these kind of plasmids for live tracking of bacteria? Where can I buy them?

Thank you.

Hello all. I purified some plasmids after doing precultures of strains from a 2-month old plate in -4 degrees. The PCR results were fine and I used theses plasmids to transform Agrobacterium cells. Should I be concerned about physiological changes for these strains from the old plate or even mutations concerning their genetic material ? (PS: I have the strains in glycerol too and I know that spread from glycerol stock should be a best practice).

Hello Everyone! I use Golden Gate technology to build plasmids in my lab. I have a plasmid that I've built that I'm now trying to maxi prep. I've already done a miniprep on this plasmid and sent it for sequencing and it was verified to be what I expected it to be. So I took this DNA and transformed it again to do a maxi prep. For both the Mini and Maxi Prep I used NEB5a cells. I've done the maxi prep on this plasmid twice and each time I'm left with just the antibiotic resistance and promoter. I have no idea where the rest of my insert is going. The GOI, Terminator, introns etc are all disappearing.

The source DNA that I'm using is sequencing correctly. I have no Idea what is happening during the maxi prep process.

Hello everyone, in the gel results image provided, the samples in lanes 2 and 5 represent linearized plasmid DNA from groups A and B respectively. Additionally, the samples in lanes 3 and 6 depict circular plasmid DNA from groups A and B respectively. Nothing was loaded to Lane 4. I am curious as to why there seems to be leakage in the sample of lane 6. Also, do you think this leakage occurred immediately after loading or during the gel run? Moreover, could the anomaly observed in lane 4 be linked to the leakage from the sample in lane 6?

I'd appreciate all your inputs.

Should I use multiple gRNA transfections (Donor) mixed as well as one by one along with a Cas9 plasmid to see which gRNA is working, or start with only 1 gRNA at a time?

Options:

Addgene options I found for all in one plasmids are pSpCas9(BB)-2A-GFP (PX458) and pX330-U6-Chimeric_BB-CBh-hSpCas9.

Which Addgene plasmids for the Cas9 would be ideal for any gene? I can clone only the gRNA sequence in the donor plasmid (Addgene) or order from a supplier. For gRNA, can I use any commercial or addgene cloned plasmids (please share a link)? What website do you believe would be the best to get the gRNA for the gene of interest?

Please help with the making this decision if you have experience with these experiments and what would be the best path to go with.

Next step is doing qPCR.

Thanks

I found a kit that they separated plasmid and genomic DNA form mammalian cell, but I want to extract both plasmid and genomic DNA manually.

I also want to use the extracted DNA for qPCR.

Anyone has the pAd∆F6 (or pAd Delta F6) helper vector plasmid map?

I did miniprep, and want to check this helper vector by restriction enzyme digestion.

Thanks,

Gang

Hi

I'm using qiagene kit to extract my plasmids DNA and recently I encountered problems to purify a lentivirus backbone plasmid (pMDLg/pRRE 8895 bp , Addgene=12251) from maxiprep.

I had no problems with miniprep. After I inoculated miniprep bacteria into big flask for large prep, bacteria grew well, every step looked fine but when I run DNA gel I found that there was only one band for uncut plasmid (looked like being linearized, but i did not add in any enzyme!) and there was smear in digested plasmid lane. To test whether my maxiprep kit is working or not, I have done maxiprep for another plasmid (pRSV-Rev 4174bp,pMD2.G 5824 bp) concurrently and it was alright for digestion of this plasmid after maxiprep, which means nothing went wrong in my maxiprep. I cannot figure out what the

problem is. I used same antibiotics (ampicilin), same LB, same maxiprep kit to prepare both constructs but I failed to produce the larger, lower yield plasmid. It is important to highlight that I have ever used the same kit to produce the lentivirus backbone plasmid for 3-4 times, I just encountered the problem of maxiprep in recent one month. So what makes my large prep plasmid being linearized/degraded during maxiprep? I have tried to pick more colonies from my amp plate and cultured with LB in small volume for miniprep and all clones are positive. I cannot understand why it cannot be purified with maxiprep kit? Anyway I hope I can solve this mystery asap with helps from you !!!

Thanks in advance :D

Here's the situation: I am currently using 6-Well plates & HEK293t cells with DMEM +10% FBS + 1% P/S and OptiMEM w/ Lipofectamine 2000 for my transfection. Before transfection when cells are 60-75% confluent, I usually change media by adding 2 mL fresh, warm media via 1 mL pipet (therefore 2x). But when doing so, cells detach very easily from the edges of the wells. Probably about 75% stay attached, but these are more localized to the center. I am transfecting in a specific plasmid at low concentration, so I am worried that this detachment will cause lower transfection efficiency in my cells and this plasmid won't get expressed due to low input. Should I redo these replicates?

I am currently trying to generate expression constructs of truncated proteins for x-ray crystallography. It is my first time doing cloning. I have had success so far in PCR amplifying my inserts using primers with NdeI, Bam HI, XhoI, and EagI restristion enzyme sites. My amplified inserts are running at the correct molecular weight as confirmed by agarose gel. I always ethanol precipitate my PCR produce overnight and resuspend the DNA pellets in 10mM Tris pH 8.0 and then do a double digest of the insert using NEB's restriction enzyme cloning tool. I typically digest my inserts for ~4-6 hours and then ethanol precipitate the digestion reaction. At the same time I do a double digest my pet28 or pet 15 vector with the same restricion enzymes under similar conditions, I have checked that each enzyme has linearized my plasmid on a gel before I add the other enzyme and then combine the digestion reactions. The uncut plasmid migrates slower than the digested plasmid and the digested plasmid runs at its expected molecular weight. I also ethanol precipitate my vector after the digestion. Finally, I use T4 DNA ligase from NEB and perform the ligation reaction. I then transform 200uL of competent cells with 10uL of the ligation reaction usually. I have tested my comp cells and used a control plasmid and was able to get good transformation efficiency with only 1ng of DNA and 50uL of comp cells but since the ligation reactions tend to give little colonies I have scaled up the transformation. I mini-prep some colonies from the transformation and have done double digests for over 100 colonies now and I do not ever see a band corresponding to my insert being there. Lately I have been doing a single digest to see if I can see the plasmid molecular weight increase as a result of the insert being there, but I do not think I see an insert also. For my most recent ligation I did a 7:1 molar ration of insert to vector in a 20uL reaction with ~20ng of vector. I have also tried 50ng of vector and 20:1 10:1, 1.2:1 ratios and still i get no insert. My most recent transformation for example gave 3 colonies for my insert reaction, 0 colonies for my control of no insert +ligase, and 5 colonies for my control reaction with no insert or ligase. I haven't digested the mini-preps yet but I feel like I will not see an insert again. I would say these results have been typical for my ligations, sometimes I get ~7-27 colonies for my reaction with the insert and 0 colonies for both of my no insert controls. When I test the colonies though, I do not see an insert or shift in molecular weight indicating the insert is not in my vector. Does anyone have any suggestions how I can get a clone successfully ?

Greetings for all of scientist using this platform. I have a little problem. Recently I had done reconstruction of my plasmid (Named PDR111, length = 11,8 kb). Transformed culture i named it W1 Transformant. After the transformation being done, I isolated the plasmid with Geneaid Presto Mini Plasmid Kit and i had done electrophoresis after i got the isolated plasmid. The results will be displayed, PDR111 is my plasmid before reconstruction (circular) as a negative control. As you can see, the band from transformed product seems to be nicked or linear. Does its mean that my transformation success? Because my supervisor told me that isolated plasmid from Presto Kit usually circular. Is it possible that my transformation product be nicked/linear plasmid? Please answer me, thank you

I'm currently trying to capture a biosynthetic gene cluster using Transformation-Associated Recombination (TAR) in yeast. After identifying positive yeast clones, I extract, via an alkaline lysis method, the plasmid from yeast and electroporate it into E. coli DH10B.

However, I have not been able to find any positive hits upon cPCR on Eci clones despite testing about 100 colonies using the same diagnostic primers I used to identify the yeast positive clones. Then, on some of the E coli that I pick and miniprep, I consistently only see my original capture vector (with no gene cluster inserted).

The issue may lie in the yeast plasmid extraction, the transformation, or the plasmid isolation prep from E coli. I know that yeast plasmid extractions are hardly ever clean and tend to be "dirty," contaminated with yeast gDNA and other DNA it has inside due to the IPA precipitation required to perform the method. That tends to lead to poor transformation efficiency into E coli. But I feel like I'm stuck going in circles trying to bring the plasmid into E coli and isolating it. if anyone has ever worked with TAR and has experienced any troubleshooting at this stage of the process, I would appreciate insight. Or any advice on things I can try to better improve yeast plasmid extraction or transformation/isolation in E coli. Many thanks!

Hi all,

I want to have 301 copies/ul for my downstream experiments. The sample I am using is a purified plasmid, of which I know the concentration (ng/ul), volume (ul), and molarity(g/mol). How can I get my desired DNA copies?

I want to copy a target gene from cDNA into a plasmid. The primers were designed according to the CDS sequence from NCBI. But when I performed PCR reactions I could not get any target bands. So the CDS sequence was synthesized into my plasmid vector. When used the plasmid as template and the above primers to run PCR, target bands were quite clear which means the primers can work. I know the gene copy number in plasmid must be much higher than in cDNA. So I increased the amount of cDNA template and cycle numbers (from 35 to 45 cycles), no target bands showed. Could anyone tell me what the problem might be. Is that possible that the CDS sequence in my cells has changed? If yes, is there any other ways to get the CDS sequence except artificial synthesis.

i need some help setting up my culture plate having negative controls,I am going to infect HEK293T cells in a 6-well plate... to one well I will add my lentivirus particles(packaged already), and to 2nd well, I will not anything and let only HEK293 T cells grow, and to the 3rd well can I add the empty lentivirus vector(having GFP and PUROMYCIN gene)? if yes,so will I add it without the packaging and envelop plasmids? and how this 3rd well can act as a control.

I am currently expressing a recombinant protein through the transduction of my plasmid into HEK 293T cells. Historically, I've encountered minimal issues and achieved high yields of protein at a satisfactory concentration. However, I've recently faced challenges as the protein expression has significantly decreased. Alongside investigating the underlying cause, I am interested in concentrating the protein obtained from my recent experiments. I am considering utilizing the Amicon Ultra for this purpose. However, I require clarification on the protocol specifics. Which buffers are recommended for this procedure? Additionally, I seek guidance on the recovery process for the concentrated protein. Your insights would be greatly appreciated. Thank you!

I tried several times using the Lipofectamine 2000 reagent, extracted the proteins but couldn't detect on Western blot. My plasmids were constructed with PcDNA 3.1+ Please does anyone have any suggestions? Thank you.

My experiment requires using blasticidin selection marker in lentivirus transduction and I did cut the sequence of interest with restriction enzyme and insert it into the backbone carrying BSR (blasticidin resistant gene). The backbone previously worked very well, and with a concentration of 10ug/mL of blasticidin, I got my stable cell line. However, this time, the transduction seemed not to work and all the cells in titration were killed by blasticidin. I first thought the concentration too high for selection. Though I switched concentration of blasticidin to 5ug/mL, no cell in transduction group could survive, which suggested something more crucial, like plasmid design, might go wrong. Here is a short sequence showing the cut sites of both plasmids for getting the insertion part and BSR backbone. Both plasmids worked very well previously.

The yellow marked region is same in both donor plasmid and BSR backbone. (including Hind III site)

The insertion part has a longer gap between its ORF and IRES. The shorter gap shows region of BSR-carrying backbone. (including Not I site)

I cut them by Hind III and Not I, and after midi prep I verified the BSR marker is inside the ligated product.

The packaging region for lentivirus is within the length limit.

Protocol for producing lentivirus was same with my previous experiment wherein same backbone was used.

Does the gap length between 1st ORF and IRES influence the expression of 2nd ORF? Though the donor plasmid can well express its selection marker?

When making a substitution to a plasmid using Q5 should I use methylated plasmid as my template? If I have a plasmid purchased from a vendor should I first transform it into E.coli?

Thank you for your suggestions. DL

Hello. I'm writing with courage to seek advice from professors and researchers. Currently, I am researching the impact of introducing a B plasmid into E. coli carrying an A plasmid, to study the effect of B genes on A genes.

After transforming the A plasmid (ampicillin resistance) via electroporation, I culture the cells to create electroporation competent cells, and then perform electroporation transformation with the B plasmid (gentamicin resistance).

While transforming A and B separately into DH10B Competent cells works well, colony formation does not occur when transforming B into A-harboring competent cells.

A plasmid: ColE1 origin, B plasmid: oriV, so there should be no incompatibility issues.

I wonder if adding ampicillin (1X, 100mg/ml) during the culturing process after transforming A could affect the cells. I tried dividing the cultures into small cultures, always adding 1x ampicillin, and when doing large cultures, I tried not adding antibiotics, or adding them at 0.2x concentration, but in all cases, transformation hardly occurs.

Should I consider anything else? Could co-transformation be the solution?

I would greatly appreciate your help. Please let me know if you need more information for your response.

Lipofectamine is a commonly used transfection reagent known for its efficiency in delivering nucleic acids into mammalian cells. However, the optimal transfection conditions can vary depending on cell type, transfection reagent, and experimental setup. Therefore, I am seeking advice from the scientific community on the specific Lipofectamine protocol that would be most effective for transfecting plasmid DNA into PC3 Eb KO cells. Any insights, recommendations, or protocols shared will greatly contribute to the success of my research project."

I am trying to clone a large (11kb) and toxic gene in an expression vector. Therefore, I am using the EPI400 CopyCutter strain that is maintaining the plasmid at a low copy number to allow bacteria growth. Attached is the protocol for this strain.

I have to grow the bacteria on plate at 28 C for 2 to 3 days to be able to get colonies.

I did a colony PCR and got 20 positive colonies.

When I put them in liquid culture (5mL to 10mL), some colonies never grown, even at 28 C. Some of them did grow (2 days incubation 28C, 250rpm in aeration culture tube, OD600 above 2.0), but then I only get a really low yield after induction of the high copy production in 100mL culture in flasks.

I tried this several times but still got a really low yield (10ng/uL, maximum 15).

I used this strain for other toxic genes and was able to get more than 40ng/uL (which is around what I need to be able to sequence my plasmid and verify the correct insertion of my gene).

I am using this kit for plasmid extraction Wizard® Plus SV Minipreps DNA Purification Systems

Any suggestions on how I could improve my plasmid yield?

Any suggestions to be able to grow the bacteria in liquid culture after getting them selected on the plate?

Thank you.

Hi everyone,

I'm in the process of creating a zebrafish Knock-in line. In order to verifying that my integration has worked, I've created a positive control plasmid with the fragment that I would expect to have in my transgenic line.

Typically, using plasmids as a positive control for PCR reactions would yield single bands due to the purity of the plasmid. My concern is that, once I optimise my PCR using the plasmid, the PCR might not actually work when using extracted gDNA from zebrafish as the template. Hence, I was wondering if it is sensible to mix the plasmid with wild type gDNA to create an unpure template. I could then use it to optimise my PCR reaction. Does this sound feasible?

Thanks :)

I am currently running a plasmid isolation. I am using a bacterial stab to grow the plasmids. We have used this stab a few times now. Unfortunately, we have not had the chance to make glycerol stocks of these plasmids. I streaked both an LB agar and LB agar with ampicillin plate. I left them in the incubator overnight at 37 degrees C. I do not see any growth or colonies. I will leave it overnight again in the incubator. I doubt that this will work because I have done this once before. If you have any suggestions please let me know. Thank you!!

I'm using the Chen&Wente protocol: https://www.addgene.org/static/cms/filer_public/02/12/0212c99c-6937-4884-8fb0-a097b965f1c3/sgrna-plasmid-construction-protocol.pdf

This is supposed to be extremely efficient, but somehow, I've never gotten it to work.  I ordered completely new enzymes because I thought my enzymes may be thawing (I re-ordered BsmBI, BlgIII, and Sal1HF), and my negative control had a greatly decreased number of colonies compared to my guide+plasmid plates for the first time; hence, I thought my enzymes were the problem.  However, upon sequencing, I still got empty vector.  Should the next step be to order new ligase? I'm not sure what's going wrong!

In 84232 add gene plasmid, I am not sure at which site should I clone guide RNA

Hello！

When constructing the plasmid, I used the same promoter. However, the direction of each expression cassette promoter in the obtained plasmid is the same, and the plasmid with the opposite direction of promoter cannot be obtained. In this case, I am worried that homologous recombination within the plasmid may occur after integration into yeast, resulting in the loss of some expression cassettes. How should I do toget the plasmid with the opposite promoter direction？

Thanks in advance！

Hi molecular biologists, I'm wondering if any of you might be able to help me with a question I have.

I am attempting to insert the DNA sequence coding for a protein domain into a plasmid (the plasmid is popinF). The insert DNA (E. coli optimised) was synthesised by Thermo (and it has passed their QA/QC), and I've successfully inserted it into popinF and transformed E. coli stellar cells, before collecting 3 different colonies from a plate to perform minipreps and acquire the plasmid with inserts. The sequencing results came back for all of them, and confirmed that the full (and correct!) DNA sequence had been inserted into one of the 3 plasmids.

However, I found it very peculiar that one of my plasmids appeared to have my DNA insert, but in a degenerated form with regards to the sequence. In the alignment shown attached, I can clearly see that there is very very strong matching of the sequenced result to the DNA from ~230 base onwards, showing that the synthetic DNA has inserted. But the sequence prior to this region does not show a high correlation to my DNA insert, and I'm wondering how this could be, and what could have caused this? I know that the synthesised DNA must be correct because I've successfully put the full length sequence into another identical plasmid - could it be that this particular plasmid showing a degenerate sequence could have undergone mutations within the E. coli or have degenerated in other ways, and if so could anybody please expand on the mechanisms and nature of these mutations? If anybody has any insight into mutation events of DNA inserts in plasmids within bacteria or knows of any good literature that reviews it and how to avoid them during recombination/transformation, I would be very appreciative for the help!

Thanks very much all,

Rob

In addgene 84323, plasmid they have empty gRNA casette. I am not sure where to clin guide RNA of interest.

1. Is it should be between U6 promoter and gRNA casette or

2. In gRNA casette itself?

I am trying to do a golden gate cloning but the ligation-digestion seems to be failing

I am cloning a PCR blunt product into pJET1.2 vector. The colony PCR and plasmid PCR have confirmed the presence of insert. The insert size is 1.17kb and the plasmid with insert looks 4kb on the gel with liberalized plasmid However, when I do digestion with HindIII and EcoRI the insert is not released. The individual digestion seems to linearize the plasmid but the double digestion does not release the insert (I tried both sequential and combined double digestion). I have not yet sequenced the plasmid.

I am performing a bacterial transformation on Mycobacterium abscessus spp. abscessus using pMSP12::mCherry. I need to know if this plasmid is integrative or replicative

Hello, I have conducted a QuikChange site saturation mutagenesis with degenerate NNS primers to generate mutant libraries of certain residues of my enzyme. For all I have used annealing temperatures of Tm - 5. The reaction mixture is the PfuUltra II Hotstart PCR Master mix: https://www.chem-agilent.com/pdf/strata/600850.pdf

Only for half of my chosen residues could I observe plasmid DNA amplification.

Generally, even if DNA amplification was successful , after transforming competent E. coli cells with the respective plasmid, only very few colonies could be seen.

What are some problems, that can occur during the PCR cycles? I dont have any major hypotheses other that primer design might need to be reassessed.

I have been trying to make a stable cell line expressing my GOI using lentiviral transduction of HEK293T cells.

To do so I ordered a lentiviral transfer plasmid from addgene which had a gene in it already. The gene is followed by an P2A sequence and then the puromycin resistance gene.

I have used this plasmid to make a lentivirus and transduce cells. This was no problem, the cells survived puromycin and expressed the protein.

But now for my GOI, I removed the present gene from the plasmid using restricition enzymes and added my GOI. Then I packaged the lentiviruses and transduced my cells. After adding puromycin, the mock transduced cells died and the transduced once survived. But, immunofluorescence staining was negative for my protein.

What puzzles me is that the puromycin resistance gene has no promotor of its own and uses the promotor and startcodon of my GOI and is expressed, but my GOI is not translated.

I checked with PCR and the gene for my GOI is present in the cells. The sequenceshow no mutations and the gene, P2A and puromycine resistance gene are all in frame.

I also tried transfecting HEK293T cells with the exact same GOI and then staining is positive.

Does anyone have a possible explanation and solution for this problem?

I am trying to select E.Coli colony which has inserted gene on pk18mob plasmid from E.Coli DH5 alpha transformation. After transformation, I did colony PCR reactions from some small colonies and it showed my inserted gene on gel electrophoresis image.

However when i cultured these colonies in LB liquid culture and extracted plasmid for restriction enzyme reaction, it did not show my inserted gene on gel electrophoresis image.

My RE protocol is 1ug of plasmid, 1ul BamHI, 1ul HindIII, 5ul buffer and water to 50ul at 37oC for 15 minutes. Then pipet 10ul to gel electrophoresis.

I obtained a plasmid from a lab down the hall. The lab has never used the plasmid before. They just happened to have it. It's labelled pMSCV-STAT6. I looked up similarly named plasmids on addgene and they're retroviral plasmids (for packaging retroviruses). However, what's confusing is the person who gave me the plasmid told me this is for packaging lentiviruses. My PI said I can't just tell a plasmid from its name. People just name plasmids randomly. So I'm really confused here. Is there a way to tell? Like if there's a particular sequence present in the plasmid that's unique to retroviral vectors or lentiviral vectors, and I can just sequence that region to determine if it's for packaging retroviruses or lentiviruses. Thanks. 

Two plasmids of different sizes were constructed based on the pBeloBAC11 backbone. One is 18kd and the other is 40kd. Choose to use TOP10 competent cells and transform in CaCl2 solution. The 18KD can be transformed successfully, but the 40kd size does not get colonies. How to optimize My method to transform a 40kd plasmid? Does the method of power transfer help?

Generally CHO (DHFR -ve) cells on transfection with plasmid bearing DHFR gene + Gene of Interest and upon addition of MTX only thoose cells which take up plasmid (containing DHFR + Gene of Interest ) will survive others will die.

My doubt 1 is : Generally DHFR is involved in De novo synthesis of Nucleotides, then how the nucleotides are synthesized in CHO (DHFR -ve ) cells?

My doubt 2 is : CHO (DHFR -ve ) cells lack DHFR so they couldn't use De novo pathway for nucleotide syntheis but they can use salvage pathway, then after transfection with Plasmid (containing DHFR + Gene of Interest) all the cells will survive due to operation of salvage pathway, now how to distinguish between the transfected cells vs Un transfected cells.

I'm confused with this DHFR-MTX selection system, could someone please help me to understand this concept, Also please share any referance material.

Hi all. I´m using calcium-phosphate transfection for 293T. I had never transfected cells with any method. I used two DNA concentrations, 5ug and 10ug per well (6-well plate). I expected lower than optimal results as it was my first time trying the procedure. I obtained about 40% efficiency with 5ug and about 10% with 10ug. My professor says that is inconsistent. I thought perhaps the DNA concentration was too high, as I seem to gather most people use 2.5ug or less, but as an undergrad I don't really know. The plasmid is TurboGFP (SHC003) and had a desirable ratio; and diluted to 5ug/ul as it was highly concentrated. 2xHBS was prepared by myself at 7.11 pH as my professor indicated; after filtering I aliquoted and froze it inmediatelly. I also prepared CaCl2 2.5M; which I also filtered and aliquoted, then kept it at 4°C.

And so I prepared:

5ug GFP: 1 ul GFP (5ug/ul), 8 ul CaCl2, 91 ul molecular grade water.

10ug GFP: 2 ul GFP (5ug/ul), 8 ul CaCl2, 90 ul molecular grade water.

I added each solution dropwise to an equal volume of 2xHBS; this was done under constant vortexing. The solution initially had a yellow-ish colour that went away quickly, and then after a couple minutes it looked white-ish.

I replaced the medium, which I had changed and hour before transfection, with 1 ml DMEM 10%SFB, 1% PenStrep.

I incubated the mixtures 10 minutes and then added it to the cells drowise and trying not to go over the same spot twice. After each drop I shook the plate gently, so the DNA could cover as much of the plate as possible. The medium turned slightly orange. I left this medium for 4 hours.

After the 4 hours I washed the cells with PBS 7 times; as I had seen very big calcium precipitates the only time I had tried it before. Then I replaced with DMEM full. I noticed while washing that some cells were lifting; even though I always take precautions not to treat them roughly.

After 24hrs I took the cells to the cytometer and got the results I already mentioned. I may have done something wrong. My professor expected at least 70% efficiency. I also added 7AAD to check for cell death/damage but I got 0%.

Any tips? Tricks? Thanks to all in advance.

I have a plasmid with kanamycine antibiotic resistant gene and Bar as a marker gene. I need to transfer this plasmid into AGL-1 strain of agrobacterium tumefaciens. I faced a problem when I follow the protocol steps of transformation. I did not get bacterial colony even after 2-days of a LB media plate having Kan 50ug/ml.

Protocol steps

1- Take competent cells from -80 C.

2- Add 5ul of plasmid having conc. 50ng/ul in 100 ul of AGL-1 BACTERIA.

3- Keep on ice for 30 min.

4- put in liquid nitrogen for 5 min

5- keep on heat bath for 5 min.

6- keep on ice for 5-min again.

7- add 800ul of LB without antibiotic (Kan)

8- Shake for 2-hrs at 28 C.

9- spread on LB media plate with kan 50ug/ml. Keep these plates on 28 C for 2-days.

But did not get the bacterial colony.

These are the protocol steps which I followed. Anyone can guide me where I am doing mistake?

In some gene editing studies using mice, the delivered plasmid carrying the editing tool (e.g. TALEN) also carries both an HA-tag and GFP. The GFP is separated from the editing tool with a T2A sequence. Why are HA and GFP both needed?

Previously I have cloned estA gene in pET28b with restriction sites EcoRI and BamHI and stored it in -80˚C. I also proceeded with site directed mutagenesis (SDM) after confirmation of the cloning via PCR gene amplification. However, I didn't get positive result post SDM. Now, as I'm trying to repeat the experiment, I'm unable to exact the plasmid from the previously cloned bacteria and stored as glycerol stock. The bacteria shows antibiotic resistance but when I try to extract plasmid and confirm through gel electrophoresis, I'm not getting any band. Please can anyone guide me and help me troubleshoot the problem.

I have plants (L. japonicus) that have been transformed with an overexpression plasmid. How can I know that these plants are homozygous for the insertion?

I noticed that a lot of plasmid annotations tend to find ColE1 origins, but I can never usually find the b. subtilis origin of replications from annotations.

I am doing plasmid isolation by Omega Plasmid DNA Minikit I #D6942-02 and Thermocycler GeneJET Plasmid Miniprep Kit #K0502. Tried both separately and no results.

I am following the instructions and using freshly prepared primary culture. All incubation periods and mixing either in the resuspension or in the lysis steps are followed carefully and I observe the viscosity of the solution after the lysis step. The white ppt. also observed after the neutralization solution. I do not know which step I might be doing wrong although my plasmid is around 6Kbp, so it does not require warming up my elution buffer. I am using sterilized distilled water as my elution. Cross checked the pipetting, chemicals, and the primary culture is really cloudy meaning that it is a rich bacterial culture. Any help?

I would to delete up to 33 bp from a plasmid to generate deletion mutation. Has anyone tried using Agilent Quikchange lightning site direct mutagenesis to delete such a long region? How feasible is this?

I think NEB Q5 site directed mutagenesis should work but this take me 2 weeks for delivery which I cannot wait for such a long time.

Any other methods will be appreciated.

Thank you.

Hi all,

I am doing a transient transfection to my cells with GFP-tagged plasmid and sending them to be FACS sorted and I need to replate them once I get them back. This is my first time ever doing this kind of experiments. would you please kindly help on how I could replate them? is it casual cell culturing? I heard that you send your plate and get a tube back?! would you be so kind to elaborate? Thank you

Hi, I ran the digested plasmid on gel and purified it separately with NEB and QIAgen gel extraction kits, but the results came like this, how should I interpret these results? Where am I going wrong or are these results normal for digested plasmid?

I have designed a genetic fusion of two separate constructs. One of them was in pET-21a and the other one was in pT7/7. Both of them expressed significantly in their respective plasmids but when I fused both of them such that one in pET-21a was tethered at 3'end followed by the other one i.e. pT7/7 and tried to express them in pET-21a, no expression was found and when I transferred my gene fusion to pET-28a, very high expression levels were monitored. I am unable to justify the reason that why is it happening as both pET-21a and pET-28a have similar sequences except the presence of his tag which only assists in purification.

After transformation in DH5 alpha, I got positive colonies. I have confirmed it by PCR (using an isolated plasmid of positive colonies as a template to run PCR by Takara Taq) and restriction digestion. In PCR, I got exactly the same size of band as my desired interest in the gene but did not get fall out of my gene in restriction digestion.

I have attached a gel pic of PCR and restriction digested . 20 ul of restriction digestion was put at different amount of plasmid ( 5 ul and 8 ul of 140 (C1 )and 305 ng/ul (C2 )

Hi. For gene deletion, I need huge quantities of highly concentrated linearized plasmid for electroporation, but, after restriction digest, I have hard time to recover satisfying quantities by ethanol or isopropanol precipitation (plasmid starting material used in restriction digest as well as linearized DNA recovered have been dosed using Qubit). Does anybody have some suggestions ?

I cloned 3HR and 5HR of p.falciparum on a plasmid and the next step is gRNA cloning. I did colony PCR for verification it is perfect but the thing is after taking the toothpeak of the colony in the petri dish should I also stream the leftover bacteria on the tip to a new plate of LB or not? is it necessary? cause the main colony already streamed once and for the second colony PCR do I need to streak it again? I will be thankful if you kindly share your experience.

We have recently aquired plasmids with the following configuration:

EF1A>{gene of interest}:P2A:Bsd

When used for transfection (293FT cells and SH-SY5Y cells), these cells expressed the desired protein after 48 hrs (verified several times by western blot and viewing of GFP which is encoded in some of our plasmids). When the selection antibiotic is added, most cells survive, which is expected. However, after a few days, the cells no longer produce the desired protein (verified many times by western blot and viewing GFP).

To be sure, we always use a negative control for the antibiotics, cells which were not transfected, and they all died quickly (36 hrs at most).

Oddly enough, when used for lentiviral infection, there is no issue, and the cells continue expressing the protein even after a few weeks of antibiotic selection.

We have not run into this problem with other vectors acquired from other sources.

Thanks in advance

Hello. I am currently attempting to select single U 87 MG cells with red fluorescence by cell sorting after transfection with Lipofectamine 3000 of a plasmid containing mCherry. The problem is that the cells do not survive after the hole process or there are few cells left that die after a few days. Does anyone have an optimized protocol for transfection and selection of U 87MG cells by cell sorting? I would appreciate.

Can I use 0.8% Agarose gel for 13.5 kb plasmid DNA?

I am co-transfecting NIH3T3 cells with two plasmids (rasV12 mutant + gene of interest) for a transformation assay. My question is: how much reagent (FuGENE HD) should I use? I typically use a 3:1 reagent:DNA ratio for single transfections. But as I am adding twice the amount of DNA in total, should I use a 3:1 ratio for only one plasmid or both plasmids? Out of situations A and B below, which would you recommend? 

Situation A: rasV12 (1 ug) + Gene X (1 ug) = 3 ul FuGENE HD

Situation B: rasV12 (1 ug) + Gene X (1 ug) = 6 ul FuGENE HD

I have gel images of the plasmid that has been digested with various restriction enzymes and an image of the transcription of digested plasmid and a plasmid map with the cut sites located. I understand that the smallest transcription product on the gel is closest to the promoter. I get a band of  transcribed RNA approx 100bp with EcoR1 so I know that on the plasmid map the promoter is either 100bp upstream or downstream of the EcoR1 cut site but how do I then know the direction of transcription. 

Hi,

I used to use lipofectamine 3000 and it worked very well. But recently my same transfections are not working (No DNA editing, while before, the same transfection was giving me 25% editing). I don't know what is the cause. The FACS analysis seems to show expression of the GFP containing plasmids in 20 to 70% of cells.

I recently noticed that my lipofectamine 3000 reagents are expired. I used one expired since 2020 and one since April 2021. But none worked.

I also noticed my optimem is slightly expired since maybe beginning 2021.

Do you know if the lipofectamine 3000 or Optimem are reagents that cannot be used after expiring date (they are both stored in the fridge at +4)

Do you have any other idea what can be the problem? I ordered new reagents anyway, so I can compare the transfections once I receive them. But I would like some opinions if people have different ideas.

I am looking to estimate the diameter (nm) of a variety of double stranded plasmids (pUC19, pMAL pIII, pKLAC2, etc.) when they are natively supercoiled and when they are relaxed.

If someone could point me towards a formula it would be much appreciated! Thanks. 

Is there any protocol of knocking out one gene in THP1 cells using CRISPR/Cas9 system ?

Here, I used LentiCRISPR-v2 system to harvest virus carring guide RNA, but when I added the virus into THP1 cells, these cells were going to be activated and differenatiated, and they would grow together.

Also, THP1 cells are a little hard to be transfected with lipo2000, so pX459 or pX458 plasmids system may be not availiable. 

So let say I have circular plasmid named "PDR111". I did PCR with KOD ONE MIX KIT and the product is linear DNA. Then, I did self circularization of linear DNA with T4 DNA Ligase protocol (Thermo Scientific #EL0014). I did everything right as the protocol said, But my transformation keep fail. Here's how my electrophoresis looks. My Plasmid size is 11871 kb. Pls guide me

Most of the GFP-ATG8 studies use yeast-derived plasmids rather than integrating into the genome. For the ones that integrate GFP-ATG8 into the genome, they are using a commercially available URA3 marker for the URA3 locus, which is not available in BY4741. Is there any other way than making a new plasmid? Thank you!

I am doing plasmid isolation by Omega Plasmid DNA Minikit I #D6942-02 and Thermocycler GeneJET Plasmid Miniprep Kit #K0502. Tried both separately and no results.

I am following the instructions and using freshly prepared primary culture. All incubation periods and mixing either in the resuspension or in the lysis steps are followed carefully and I observe the viscosity of the solution after the lysis step. The white ppt. also observed after the neutralization solution. I do not know which step I might be doing wrong although my plasmid is around 6Kbp, so it does not require warming up my elution buffer. I am using sterilized distilled water as my elution. Cross checked the pipetting, chemicals, and the primary culture is really cloudy meaning that it is a rich bacterial culture. Any help?

Hi, I' am trying to make stable cell line using plasmid transfection.

the plasmid are available for neomycin selection.

I want to know the appropriate G418 concentration and time forselection about Mlg cell line. Mlg cell line is mouse lung fibroblast cell line.

and also I wonder the concentration and time of selection about mouse primary lung fibroblast.

Thank you