Plasmids - Science topic

The material at the bottom of the gel could be degraded DNA or it very likely is just tRNA and other stable RNA that are in your plasmid prep.

Thank you for help, I repeated the experiment and got positive results.

see this:

Hello. It is better to use the TA cloning method for cloning large pieces. You can also make changes in your ligation to increase the efficiency of the reaction. You can use the site https://nebiocalculator.neb.com/#!/ligation to get the exact values of the vector and insert, in the ligation reaction. Also, changing the strains of bacteria used in transformation or their preparation method can help you. TOP10 bacteria is a good option for preparing competent bacteria using calcium chloride.

Thank you so much Gina Lauffer for your brief response. Actually I missed to elaborate that when I say plasmid, it means plasmid vector, as these vectors are used in gene therapy to cure sevral diseases. so when these vectors are present inside eukaryotic cells (eg human), how does histone present in cytoplasm of eukaryotic cells bind to these vectors.

Could you please help me gain a better understanding of this?

Luigi Marongiu how about trying plasmidSpades? If i understand your question correctly.

Based on your description, it sounds like the transfected cells are experiencing cell cycle arrest or apoptosis due to toxicity from the transfection reagents and/or overexpression of the transfected plasmid.

You can try to:

- Reducing the amount of transfection reagent even further. Start with the manufacturer's recommended minimum and titrate down. Too much lipid can be quite toxic.

- Optimize the ratio of DNA:lipid. Test different ratios to find the sweet spot for your cell type.

- Try a different transfection reagent. Lipofectamine is quite harsh. You could try FuGENE or Polyjet which may be less toxic.

- Reduce the amount of plasmid DNA transfected. Overexpression can trigger apoptosis. Titrate down to find the minimum needed.

- Use a lower CMV or inducible promoter to reduce expression levels.

- Try transfecting plasmid expressing an inert reporter like GFP first to rule out plasmid toxicity.

- Let cells recover overnight in normal media before imaging.

- Check for apoptosis markers like cleaved caspase-3 in transfected cells.

- Monitor proliferation with labels like EdU or by quantifying cell numbers over time.

With some optimization of reagents and DNA amounts, you should be able to find conditions where transfected cells still divide.

The concentrations are pretty decent, how much are you wanting? The contamination issue can be helped by an additional wash step. Also, if it's a column, try to avoid transferring over any cell debris after lysis and clearing.

Yes, it's very common to use two different antibiotic resistance selection markers in mammalian cell culture. However, 293T cells express a neomycin resistance cassette, and this confers resistance to G418 (also known as Geneticin), so you cannot use G418 selection in 293T cells.

I think that the logic is that 1 unit of RE is defined by the time that it takes to cut a defined amount of dna. This dna is often human dna. If the enzyme is a 6 cutter and if we assume that the distribution of bases in human dna is random then a 6 cutter cuts every 4 exp6 or every 4096 bases but in your pcr product which is 500bp there may be one cut site in every molecule so the enzyme is having to cut 8 times as many cut sites for any given amount of dna so it is wise to give the enzyme longer so that it can cut all of the RE sites. In genomic dna partials are not very obvious but partial digests of a linear pcr product just look uncut which may lead you to think that the molecule has not cut therefore the cut site does not exist so you will charaterise the pcr product as having the wrong genotype

You can also use recombinases for site specific integration or transposases for random insertion to incorporate your sequence into the genome.

If you really, really want to know, then sequence the plasmid.

But since you have several colonies that are exactly what you expect, just choose those ones and continue with the experiment.

Good luck!

Dear Mohamed Samir Youssef did you try to pate your Agro without transformation or a mock transformation to see if there is some contamination that is not Agro or an Agro resistant to your antibiotic of choice or that antibiotics are ok? After purifying plasmid from Agro, do you run it on gel before doing PCR? normally the concentration is very low, but you could still try to see if you got any plasmid at all or nothing.

Hi Huang,

Lipofectamine 3000 reagent certainly worth trying if you're facing low transfection efficiency with HEp-2 cells. Always perform control experiments and assess cell viability and transfection efficiency to ensure that the new reagent is working effectively for your specific application. Additionally, consider running parallel experiments with your previous reagents for comparison.

Transfection can be a trial-and-error process, and the right reagent and conditions can vary depending on the cell type and the specific plasmid being used.

Barely. You can visualize a single band that has maybe 5-10ng in it, although faintly. So you should be able to see that amount of DNA if in a single band. But after ligation you might have a number of various ligation products so that each band could be faint.

I never bothered running ligation products on a gel, I just ensured that I ran the proper controls and then transformed.

Thank you so much. I shall introduce an extra wash step and check Michael J. Benedik

Hello. I got the same issue when doing transfection on human cardiomyocytes. I just wonder if you could find a solution for that issue. And one more question is how could you standardize the amount of all plasmids for the transfection, by mass or by copy number (molarity).

Thanks Yoram Gerchman ! Very good points.

I am not sure about your protocol, but in these kind of transfections, optimising (reducing) the DNA amounts, PEI amounts and doing transfection under antibiotic free conditions will be required to get optimum viability/transfection. Best!

Heating does not remove supercoiling. It requires a nick or break in the DNA by an endonuclease or a topoisomerase or to remove it.

It should be fine, I would not expect 3 days at -20 to be an issue.

Bruno Salomone Gonzalez de Castejon gave an excellent answer, there are many instances of multiple plasmids in a bacterium, even with natural plasmids in populations from the wild.

I think it’s too much concentration, You should try to dilute the cells.

Dear Yukti

you can certanilly do it trough mutagenesis. However i suggest to you to evaluate enzime free cloning approaches as the PIPE cloning (that could be also used for plasmid mutagenesis) that let you free to the presence of restriction enzimes.

if you are interested to know more details about it, read the following papers

or look to the following videos available on my blog, ProteoCool

best regards

Manuele

There are a couple of controls you can run to try and figure out how much uncut or cut and self-ligated plasmid you got. Here is an example:'

tube 1: cut and purified vector, ligase buffer, water, NO insert, NO ligase

tube 2: cut and purified vector, ligase buffer, water, NO insert, with ligase

tube 3: cut and purified vector, ligase buffer, water, your insert, with ligase

tube 1 will let you know how much vector was uncut

tube 2 will let you know how much vector was uncut or was single cut and self-ligated

tube 3 is your actual ligation reaction.

Ideally, tube 1 and 2 have 0 to very few colonies, and tube 3 has more.

In addition to what Robert Adolf Brinzer writes, adding some carrier such as tRNA helps to precipitate your DNA if it is at low levels.

Regarding using ethanol, it works but you need a final concentration of around 75%, in other words adding 3 volumes of 100% ethanol to your solution.

The easiest solution would be to use a serine/threonine recombinase like CRE or FLP to integrate your plasmid into the genome. If you want a specific insertion site use CRISPR to make a double strand break and then insert a landing site with selectable markers between the two homology arms. With recombination sites at either end of the selection cassette you can then do cassette exchange. Using a lethal gene driven by a tet operated promoter can allow for selection of your fragment being inserted into the genome.

For getting your fragment I advise checking if there is a cosmid library with your region of interest. Else you may need to clone in fragments and then do a Gibson assembly.

Try giving it a few more days. Sometimes you'll need 7-10 to see some growth. Also, try making single plasmid selection plates to check that both your bait and prey plasmids are giving good transformation and growth on their own.

Those ITR seem to be Inverted Terminal Repeats sequences, which are used to package the expression casette on adenoviral vectors: https://blog.addgene.org/viral-vectors-101-parts-of-the-aav-transfer-plasmid

Thank you!

It seems very likely that your strain is contaminated with another plasmid, a 3kb plasmid is about the right size for a pUC or pBK type plasmid. The copy number of large plasmids is often much lower so it is possible that you just can't see it if it were there. But I see hints of a band around 25 kb range in the undigested samples. The digest lanes have that strange reflection making it impossible to see.

Most bacterial strains should be able to support larger plasmids. Note that 36 kb isn't really that big, many natural plasmids are larger than 100kb. But it could depend upon the type of plasmid etc. But if it works in DH5alpha it will probably be fine in nearly any E coli strain.

You could PCR your insert and add a different set of restriction sites on the PCR primers so that you can clone it in reverse orientation.

I would also suggest if you are having f1 phage contamination issues, you switch to a host strain that is not carrying an F episome. Filamentous phages like f1 can not infect a strain that is not carrying the F episome.

The answer is going to depend upon some various things.

First of all, when you write BL21 do you really mean BL21 or do you mean BL21(DE3). The two strains are different. Many people use BL21 as shorthand for the DE3 strain, but it is in fact different.

Secondly, what is your expression vector, or more precisely what promoter is being used. BL21(DE3) is designed to permit expression from plasmids with the T7 promoter, such as the pETxx series of plasmids. If you are using one of those plasmids, then it will not work in HB101 and you need to use BL21(DE3) or something similar that provides T7 polymerase.

On the other hand if you are using a plasmid with a different promoter, then you probably don't want to use BL21(DE3) because expressing the T7 polymerase can have impacts on the cellular physiology.

Additionally HB101 does not adequately regulate any promoters based on lac, in other words any promoters that require IPTG induction. Generally HB101 was designed as a cloning strain and not an expression strain, although it can be used for expression in some circumstances.

So the choice of strains depends upon the expression plasmid, the mode of regulation of that plasmid and to some degree the protein you are trying to express. If you provide more specifics you might get a more detailed answer.

I agree with the nanodrop detection threshold being problematic.

Another item to consider is copy number going into each qPCR reaction. Your QPCR machine will have upper and lower limits, if you get out of that range then the data won't be linear for your curve.

The uniformity can often be improved by making "master mixes" for any given sample concentration then aliquoting into the triplicate wells.

Thank you so much for the recommendation. Wish me luck!

Hi again, just for info you don't need large sequences of homology to promote recombination. 20 to 40 bases are enough...

In such scenarios, it would be beneficial to pause and conduct some quality checks.

Thank you for sharing the image of your cells pre-transfection. However, could you kindly provide pictures of the mock control as well?

Moreover, I would suggest including an additional fluorescent control to demonstrate the efficacy of transfection.

This data will assist us in comprehensively examining the situation and evaluating any potential cytotoxicity concerns.

Best wishes!

Prasad Trivedi Thank you for suggestions!

Michael J. Benedik Thank you so much professor for your help.

I guess you just need to try it and find out. Most likely it will be fine.

As mentioned Wizard® Plus SV Minipreps DNA Purification kit from Promega is one of the best kits for good quality AND quantity plasmid isolation with a close competition from the NEB's Monarch kits and favorogens plasmid miniprep kits. And also as mentioned above use an endotoxin removal based kit for your work.

Kits vary with their plasmid recovery based on mainly the silica bead packing, the thicker bed columns do have the potential to bind a lot of DNA but they require a lot more elution volume as well and the plasmid gets highly diluted. Thus, columns with thinner beds in my experience seem to give best results.

One strange thing I have also observed is that when I left columns at 4 degrees for some time and then used them they gave me a better yield, I am unaware of the reason or whether this phenomenon is actually true or not but since it seemed to work for me I can suggest this as well.

One final thing incubating the bacterial cultures on ice for 1-2 hrs before plasmid isolation seems to improve the Superhelical plasmid yield as well. Maybe because the DNA repair/ packing mechanisms getting activated during the dormant growth stage in bacteria.

Thank you Robert Adolf Brinzer sir for your valuable response.

How did the cell cultures look after the incubation? Nice and cloudy? If there are few cells going in then you'll get a low yield. Is anyone else having the same problem using the same kit? As mentioned above, kits do expire. It can help to warm the elution buffer (50C) then let it incubate for a minute on your column prior to eluting.

Since you are not wanting to keep the backbone as an extrachromosomal element you don't care if the origin works in the destination species. Yes you want a bacterial antibiotic selection marker for convenience when cloning. Yes you ideally want a selective marker for your target species in your insertion cassette. The main factor for choosing which backbone to use is a small size to increase transfection efficiency and the restriction sites in the MCS which must be compatible with restriction sites in the homology arms. Adding a single strand annealing protein expression cassette outside your insertion cassette can increase homologous recombination frequency, improving the likelihood your knock in occurs. This means you can use pUC backbones, pBluescript or whatever you prefer.

The choice to subclone often has to do with how precious or heterogenous your template samples are. Have you run your restriction digests for cloning on a gel? Are your restriction enzymes still before their expiry date? Sometimes performing an overnight digestion can help. Are you using a CIP treatment step or not?

Robert Adolf Brinzer Thank you very much

The answer is yes but like all transfections the larger the plasmid is the lower the transfection rate.

Lipofectamine is a mix of branched polyamines and modified fatty acids which encapsulate your DNA and allow it to fuse with the cell membrane. More recent iterations also have endosomal escape functionality. This means that unlike viral transfection there cargo size is not limiting.

What auxotrophies are you using?

Generally you don't need sterile evironment for DNA extraction.

Sam Carter try electroporation definitely. Heat shock is finicky. Use DH10B strain E. Coli. thermo Fischer sells them

If you are gel extracting your band then usually a contaminating plasmid is not an issue. But yes the DNA is likely to always be present. If this is an issue, another solution would be to do a mini prep and then retransform some clean cells and select only for your Amp plasmid. Most of the transformants will only pick up the one plasmid (but you should test and confirm before proceeding).

It would appear that you have an insert of around 450nts cloned into your plasmid, based on the simplest explanation of the figure you attach. Does this sound like what you should have?

If the insert is correct then of course you can make a probe from it by releasing with appropriate restriction enzymes or by PCR amplifying the insert region.

Emma Thompson 6000bp and STBL3 and TOP10F

I am assuming when you say "it is not happening" means you are unable to see the 400bp insert in the gel? You do not say how much plasmid you digested but more than likely there is not enough insert in the gel to visualize.

Your 400bp insert is ~12% of the overall 3400bp size of the plasmid so if you digested only 100ng of plasmid, you would only have ~12ng of insert to see in the gel which is on the low end of what is possible to see in an agarose gel. If you double digest 500ng of the plasmid you should be able to easily see the insert (~60ng).

Lauren Woodward Hartzler

Competent Agrobacterium cells:

Transformation:

Robert Adolf Brinzer Thank you for answering my question.

This is experimental evolution and 10 μL culture was transfered to 980 μL LB medium with 10 μL low-dose antibiotics every 24 hours. The ratio of cells carrying the plasmids was identified every 6 days. Fluorescence was monitored by the Beckman CytoFLEX SRT. Overnight cultures were diluted 1:100 with PBS in 5 ml culture tubes for loading on the flow cytometer. The channel of emission detector wes set as P1: 561-Yellow (Y610/20 nm) for PE-mCheery. For each sample, 30,000 cells were collected. And the gain for mCheery was 1000.

Hi Sweta,

We recently published a paper that well answer your concerns and offer a nearly perfect tool for stable overexpression of lncRNAs. In this study, we found that the lentiviral vector produces lncRNAs with improper termination, appending an extra fragment of ~2 kb to the 3’-end. Consequently, the secondary structures were changed, the RNA-protein interactions were blocked, and the functions were impaired in certain lncRNAs. To solve this problem, we developed a novel lncRNA expression tool called Expression of LncRNAs with Endogenous Characteristics using Transposon System (ELECTS). By inserting termination signal after the lncRNA sequence, ELECTS produces transcripts without 3'-flanking sequences and retains the native features and function of lncRNAs, which cannot be achieved by lentiviral vectors. Moreover, there is no potential risk of infection for the operators and it takes much less time. ELECTS provides a reliable, convenient, safe, efficient and time-saving delivery tool for stable expressing lncRNAs.

The open accession of the full article is available at : https://jnanobiotechnology.biomedcentral.com/articles/10.1186/s12951-021-01044-7

We welcome all the researchers to use our tools. If you have any questions, please feel free to contact me, my email is: zhangy525@mail.sysu.edu.cn.

some mammalian sequences is hardly replicated in E. coli. Also, large plasmids makes replication difficult. Try to use E. coli strain appropriated for replicating mammalias sequences such as XL10 gold.

Martin O'Reilly

thank you for your reply.

our lab just have BTX electroporator.

I would consider checking if they have a f1/fd/m13 ori annotated.

Maybe, but you got so few colonies with the positive control that if the quality of your recombinant plasmid is not as good, you might just not be getting anything.

Have you checked the quality of the plasmid? You should only need to use a few ng (not 1000 ng) for a transformation.

Marzieh Rostaminejad Thank you. pCMV-3Tag-4A and pEGFP-C1 for this vector which one will be expressed ? I saw the gene map but I am not sure which one is functional ? Can you explain it.

If your CRISPR vector is lentiviral based, I think it is better to use other strains of component cells instead of DH5a. It is easier to acquire undesired plasmids in this scenario.

Huyen Ngo Please, repeat both PCR in small and large volume, and if small is OK, just amplify DNA in several tubes for sequencing. Large volume has different kinetics of the PCR.

PCR by site directed mutagenesis produces a nicked circular PCR product that hasn't been ligated together into a plasmid. Transformation to E.coli cells ligates the PCR product together using native E.coli DNA ligase and ATP. However, before that happens occassionally you get really unlucky and the excess of primers from the reaction gets inserted into the nicks and then the plasmid gets ligated together with the primers inserted. . Use of perfectly complementary site directed mutagenesis is not actually PCR. There is no chain reaction as the primers cannot extend over the nick. So the QuickChange site directed mutagenesis doesn't work so well. It is just linear not exponential amplification.

Use of partially overlapping primers creates a nicked circular PCR product. The PCR product remains nicked all-throughout but the primers anneal over the nick and amplify the PCR product so it is a true chain reaction amplification. This method is more likely to work because it is actually PCR, but also more susceptible to primer insertion events.

I am not quite sure but looks like either the forward primer or the reverse primer or their complement was inserted like 5 times or a mixture of all the above. One thing you can do is translate the amino acid sequence of this insertion to see what the pattern is. If the insertion is in frame your enzyme could still be active with the insertion folded away from the rest of the protein. Alphafold it, but you should probably throw away this plasmid. It would be too hard to explain even if it was active.

One thing you can do to make this happen less is to use a Qiagen PCR product purification kit to remove the primers from your PCR reaction product before transformation to E.coli cells. This makes the primer insertions less likely to happen. but it still happens to me.

Also, Sanger Sequencing of plasmids is kind of obsolete. Oxford Nanopore sequencing is really good. Is that what you are using? You can use Plasmidsaurus. Just ship them your plasmid and they give you results back in like 2 days for the whole plasmid using nanopore sequencing.

Did you measure the protein concentrations of the extracts used to prepare the samples you ran on the gel?

If you are not getting that particular plasmid circularized while others were circularized, you should get an idea what could be the problem. Either there are some parts missing or there might be the quality issues. Thus, these issues are probably not easy to fix based on the data. If you are changinf the paramters, you might accomplish your task but there might be a trade off on the quality.

Using more template just means there is a slightly higher chance of transfecting your starting plasmid into your competent cells and not your PCR mutagenized product.

i was having the same doubt being the limit packaging size i found for lentivirus is 9Kb.....did you find any answer to this question?

I am not sure about the answer. But according to your concern, maybe it is better to use inducible system (ex: tet on) to express the fluorescent protein?

If you think you may have M13 contamination, then transform your plasmids into a non-male host strain (F-) which won't be infected by M13 and then make a new DNA prep.