Plasmid Cloning - Science method

Check the Alphafold predicted structure of your protein. If the N-terimus is deeply buried in the protein or involved in protein-protein interactions, try the C-terminus. Probably best to do both though since it isn't that much extra work.

Has anyone tried GenScript's GenBuilder DNA assembly kits?

Hi, DH5alpha is not affected by salt concentration, so either formulation is fine.

From my experience, the low salt LB is used for work with antibiotics zeocin, which is salt-sensitive. Of course you could have also some osmotic-sensitive strains of E. coli, but that's not the case of DH5alpha.

How big a size difference are you talking about? If you are talking about a few kb larger then I would not expect a lot of difference, but if say double or triple size then quite possible. Another thing to consider though is efficiency of plasmid isolation. As you get to larger plasmid sizes you may shear more of the plasmid or just not successfully isolate it during the purification process. Lastly it might have nothing to do with size but with the insert.

Well, creating a semi-permeable transcription bubble can be challenging in the context of the structural stability of DNA as it tends to reanneal to its double-stranded form. Next is the concern of replication, which involves the fact that semi-permanent unwinding can potentially hinder the replication machinery from proceeding with DNA replication. Lastly, to maintain the transcription bubble to its semi-permanent unwound state, RNA polymerase is required to be halted in its activity at the transcription site, which could, in turn, lead to instability and interference in the replication of the plasmid. Considering these aspects, I believe three probable yet theoretical strategies can be adopted in this regard. First is genetically engineering a modified RNA polymerase, which can maintain the plasmid DNA at its single-stranded state by getting associated at a precise plasmid location without hindering the transcription process. Second is implementing genetically engineered single-strand binding (SSB) proteins, which can keep the plasmid DNA at its unwound state without interfering with RNA synthesis. Lastly, chemical molecules such as intercalating agents are introduced, which can develop proximal unwinding by being inserted at the nitrogenous base pairs of plasmid DNA; Molecules that are enhancers or activators of helicases; Hydrogen bond destabilizers like Di-Methyl Sulfoxide (DMSO), Urea or Formamide which can perform denaturation of double-stranded DNA; Cross-linking agents like Psoralens which forms covalent cross-linkages between single-stranded DNA molecules and DNA or RNA polymerases; Ligands which associate with single-stranded DNA such as Peptide Nucleic Acids (PNAs) and nucleic analogs which stabilizes the single-stranded structures of DNA; Alkylating agents such as Nitrogen and Sulfur derivatives of Mustard gas, Ethyl Methanesulfonate (EMS), Methyl Methanesulfonate (MMS), N-Nitrosoureas and Temozolomide. Nevertheless, besides being hypothetical, all these strategies have cons of their own.

Dear Colleague,

I hope this message finds you well. Cloning a custom shRNA sequence into the pGIPZ plasmid involves several meticulous steps to ensure success. Here, I provide a detailed protocol to guide you through the process:

By following these steps meticulously, you can successfully clone and express a custom shRNA sequence in the pGIPZ plasmid, enabling efficient gene knockdown in your target cells.

Should you have any further questions or require additional assistance, please feel free to reach out.

Reviewing the protocols listed here may offer further guidance in addressing this issue

Dear S. Sharif

I am struggling with a similar problem, my untransformed cells are growing on Zeocin plates. How you manage to solve the problem?

Thank you

These are sephadex spin columns and work on the basis of size exclusion. So dna of more than 10 bases long will flow round the beads and elute early while small salts will have to flow through the pores of the beads and will elute much later. They can be used for dna purification from most smaller molecules and have been used to remove radioactive salts from end labelling of oligos with the labelled oligo eluting first off the column. The G25 is an indication of the size range that can be separated on these columns

Try precipitating your plasmid with LiCl and ethanol and use less water to resuspend it. Scaling up is for more ug of DNA

If your question is whether this plasmid will work well in plants, the answer is that it will not. The above plasmid is designed as an E. coli expression vector and only for certain strains of E. coli. The plasmid will not replicate in plants nor will the promoters function unless you also provide the T7 RNA polymerase.

Jeremy Mullesa I think your plasmid is ok. PDR111 is overloaded, therefore, it appears to run faster and looks smaller/different from your clone. Load equal amounts. To ensure you have the correct fragment cloned, sequence your construct.

Kais Khudhair al Hadrawi you answer is off-topic and simply generated by ChatGPT. The RG community for sure knows how to use this tool.

When conducting experiments involving the lentiCRISPR v2 system for CRISPR-Cas9-mediated gene editing, verifying the correct insertion of the guide RNA (gRNA) sequence is a critical step. This ensures the specificity and efficacy of the gene editing process. To facilitate this, PCR amplification of the gRNA cassette followed by sequencing is commonly performed. Utilizing specific primers that flank the gRNA cloning site within the lentiCRISPR v2 vector is essential for this verification process.

Primer Design for gRNA Insert Verification:

The lentiCRISPR v2 vector, a widely used tool in gene editing, incorporates a U6 promoter driving the expression of the gRNA, followed by a cloning site where the gRNA is inserted. To amplify this region and confirm the presence and correct orientation of the gRNA insert, primers that anneal to sequences flanking the cloning site are used.

Suggested Primers:

PCR Amplification and Verification:

Conclusion:

The use of well-designed primers specific to the U6 promoter and the region downstream of the gRNA cloning site is essential for verifying the correct insertion of the gRNA in the lentiCRISPR v2 vector. This verification step is crucial for the success of CRISPR-Cas9 gene editing experiments, as it directly impacts the specificity and efficacy of the editing process. By following these guidelines for primer selection, PCR amplification, and sequencing, researchers can confidently validate the presence and accuracy of their gRNA inserts within the lentiCRISPR v2 system.

Take a look at this protocol list; it could assist in understanding and solving the problem.

In addition to the previous responses, if you rule out primer contamination I also suggest trying to re-isolate the positive colony. Sometimes we think we got only one colony, but what looks like a single colony is a merge of two very close colonies or two overlapping colonies. You can try re-streaking your positive clones and screening colonies from that. This will help explain what's going on.

You may also have both the empty vector and the plasmid of interest in the same bacteria (bad luck!). If that is the case you should probably start over the transformation. If you move on with it, your cells may end up spitting the plasmid and keeping the empty vector for antibiotic resistance (since the empty vector is small and less energetically costly for the bacteria to maintain).

But remember it might be the original measurement that was off. Lastly it might not matter, use the lower concentration as real. and It won’t hurt if there is a bit more than expected Unless you need to be very quantitative

Plasmid recombination following transformation can be a significant issue, especially when working with constructs that have repetitive sequences, large plasmids, or multiple plasmids being transformed into the same cell. Several factors can contribute to plasmid recombination:

1. Presence of Repetitive Sequences

2. Plasmid Size and Complexity

3. Host Strain Recombination Activity

4. Transformation Method

5. Multiple Plasmids in One Cell

Strategies to Minimize Recombination:

Addressing these factors and implementing strategies to minimize their impact can significantly reduce the occurrence of unwanted recombination events during plasmid transformation.

l Take a look at this protocol list; it could assist in understanding and solving the problem.

if the two parts are fundamental for your gene , please purify the bands from the gel , use the ligase and add adaptors for a new cloning by PCR, then you can re-clone using new enzymes fit for the adaptors

Daniela Liebsch Thanx for wonderful explaination

Excuse me, has someone got the answer？

Robert Adolf Brinzer My siRNA segments will be 21nts long, T- stretch for terminantion will be 6 T long.

First of all, I don't think ZYCY10P3S2T is RecA-, at least the one listing of the genotype did not show that it was.

Secondly, when you run uncut plasmid, most commonly you see a mixture of supercoiled and nicked plasmid, which will have differing mobilities. So a pure monomer prep will still normally show two bands from uncut plasmid that resolve into one band.

It seems to me that you are not seeing mulitmers, merely nicked and unnicked circle.

I also had this problem. Before you run the gel, purify the ligation product to remove the buffer and the T4 ligase, then you will see the band.

I would also be interested in the sequence of pLO1.

Do you know where I can get the plasmid? Can I isolate it or somethink like this?

Dear Carolina

I also faced a similar problem. I cloned the 5´-UTR of a gene above the luciferase sequence of the pGL3 control vector, but after inserting, the amount of luciferase decreased drastically, while the frame was not disturbed. I guessed that the presence of an ATG in the inserted sequence is the cause of the problem. Therefore, after removing this ATG, I observed that the expression of luciferase returned to its normal state. I suggest you change or delete the ATG upstream of the luciferase sequence (especially if ATG is the beginning of the first exon.)

Best regards

if your plasmid has no large repeat (like LTR from retrovirus) you can certainly transform dh5... the stbl are used for unstable plasmid that can recombine... just check that after purification your plasmid is still the same...

Ask your professor for help with your homework. This site is for research projects, not cheating on assignments.

Plasmids for mammalian expression use different organism specific promoters, a eukaryotic ribosomal binding site, an intron in the CDS of the gene of interest to avoid bacterial expression and to increase expression in the mammalian cells and a poly A tail after the stop codon to reduce mRNA degradation.

Simply add low material

So I have successfully cloned shRNA into this plasmid. I used T4-PNK (NEB) alongwith 1ul of NEB ligase buffer to charge phosphate groups at the end of the oligos. This was done in PCR machine (37 deg for 45min and then 95deg for 5min then -1deg/20sec), 10ul rxn vol..

I know that double digest has the phosphate groups to work with but this has helped me a lot.

Then dilute the product to 30-50 times. Use 1ul to 2ul (You can use more but this T4 PNK charging will give you the result), 1ul (27ng) of double digested DNA and 1ul T4 DNA ligase (I used Takara but it hardly matters) and obviously buffer (Final volume of the reaction is 10ul). Then use 3 conditions (37deg, 4h; 16deg, 16h and 4deg, 48-72h). I have used DH5A and NEB Turbo (Ultracompetent cells), in NEB turbo you will get a ton of colonies but that will add up to your background as well. I got +ve colony from DH5A. I have screened 40 colonies and got positive in one of them. I had tried colony pcr but it gave me false positive. So RE digestion seemed to be more promising. Please note that I did 80 mini preps, without using kits, in 2 days and I was able to do that in 2ml MCTs with 1ml culture volume, you will get enough DNA to do screening (anyway you need 1-2ug for screening). I used 0.15ul of enzyme, XhoI, as it is pointless to spend much on enzyme if you want to see just 2 bands at 200bp. During minipreps I used RNase and PCI, but without these also you should get result.

Hi, Ruojin Tian

I do agree with all points commented by Mimi Asogwa and would just make a few questions to figure out what is happening.

1 - Why did your PCR product did not give you a one amplicon only? Did you try everything to optimize it or just followed some paper protocol? If you just followed someone else's protocol, then, optimize it and then use it. Not optimized reaction may lead to this kind of problem.

2 - Usually in all assembling reactions, again as Mimi Asogwa told you, it's better to use/try different vector/insert ratios and use a molecular calculator to find the most precise mass amount (in nanograms) for this purpose. So, a 1:1 ratio is not always the best (honestly, only when the sizes are very close to each other).

3 - Lastly, you could do it stepwise. First, take the fragments that makes your complete insert together and only then, a second reaction with it and the vector. Sometimes, it works better.

Hope my comments may help you.

The direct repeats tend to not be stable in E.coli during cloning. I would suggest you simply clone your gene into two different integrative vectors (would integrate in two different loci) then you can have two copies of the your gene. If you really need the same vector you can try two different strong promoters to reduce the homology and see if that works.

Noted

Well, the comment about the His-tag does not really answer the question about nuclear import efficiency?! I think it is not fully predictable... some proteins are very well imported with a single SV40-NLS. For others, import may be improved by adding another copy. We worked with Cas9; import improved a lot with a 2nd NLS (see Gruetzner et al 2021 Plant Comms). Some Cas9 versions have 3 - 5 NLSs attached, still no negative effects. If you are worried about using 2xSV40, just use two different ones. I made good experiences with the c-myc NLS in plants, we used ATGGCTCCTGCTGCCAAAAGAGTTAAACTCGACTCAgctgccgcA fro N-ter fusions, tctgcaCCTGCTGCCAAAAGAGTTAAACTCGACTAG for C-ter fusions including some linker amino acids. The core motif is PAAKRVKLD.

difficult to find a good ER site for the cloning.

difficult to clone (pcr) the whole gene at once, need to find ER site to link/ligate it.

a large fragment is difficult to ligate and transform.

not all the vectors can work well with large fragments.

etc.

Thank you for your response. I will check upon it.

In my opinion, this effect is not attributable to usage of the pCDNA3 vector or cmv promoter. This may be due to the sequence included by the experimentator. Maybe, it contains a weak termination signal/sequence or additional spilcing sites leading to the generation of a truncated protein additional to the full lengh form.

More likely is that the antibodies used for detection of the resulting protein via WB is not really specific and could possibly detect additional proteins together with or instead of the protein of interest (for ref. see Baker M. Reproducibility crisis Blame it on the antibodies. Nature. 2015_521.p274-6. doi 10.1038521274a).

Electroporation has been used successfully to deliver plasmid DNA to a variety of tissues in vivo. Because of its physical nature, EP can be applied to practically any cell or tissue. Plasmid DNA in the appropriate diluent is injected into the tissue. Electrodes are then placed around the injection site and the cells within the tissue are subjected to a high-voltage electrical pulse of defined magnitude and length. The animals are then allowed to recover and the tissue is evaluated at specified time points following delivery. Factors that can be varied to optimize electroporation effectiveness are pulse width, number, amplitude and electrode configuration.

It may be that your original fragment before purification has significant secondary structure and is supercoiled and knotted to a small spherical shape. In DNA purification kits the sample to be purified is added to chaotrophic salts which linearise the molecule to facilitate the binding of the dna to the membrane.If the eluted dna remains linear on elution from the membrane then it will run slower through the agarose and will appear as a larger size than the natural coiled version

Yes, you should remove the RNase from your sample.

Paul Weber Maybe your RNase might be the issue. I have never got RNA contamination using Qiagen Miniprep Kits. Please, recheck if you have added RNase provided in the kit to P1 solution, if you have added it then RNase might be faulty. You can add more RNase to P1 or go for a separate RNase treatment step after your plasmid extraction. All the very best for your experiments.

That is a shame. Not1 cuts well in NEB buffer 3.1 or cutsmart buffer but badly in other buffers ( only 10% activity in buffer 1.1) while Xho1 cuts well in all salt concentrations and buffers so if you had cut with Not1 first then you could either have done a double digest or a second digest using Xho1 without purification but cutting with Not1 second is much less forgiving. You can purify your partially double cut product and re digest with only Not1 since it has cut already with Xho1 if you have enough to digest and re purify and still have enough for the next stage of your experiment

Hi,

bacteria can have the gene orientation on one and the same strand of DNA in different directions. So, when you look at a software showing you the strand it can only go in one direction. This does not mean that antisense strand and sense strand are defined it is always related to the gene you look at. Your gene is always encoded by the coding strand.

Maybe try imagining turning your double strand, which you are looking at, by 180 degree and read the gene from 5' to 3'. Double check if the first three amino acids encode Met.

I can imagine how one can get confused about it, so its a bit tricky not confusing you even more with my words.

When you are using a software, the plasmid should be also displayed from 5' to 3' (the upper stand is always the coding strand which goes from 5' to 3', in case it displays the double stand)

Your gene should come after the promotor. You connect the 5' of your coding strand to the 3' of your linearized plasmid (copy past it in the gap where you want it to be). When you are done with insertion the whole sequence should be readable in 5' - 3'.

Wish you all the best ;)

Hi. There are several advantages and even more if you aim to assemble several BioBricks.

I found this nice explanation at addgene blog : hope it helps you to clarify :

"Advantages of Golden Gate cloning

Golden Gate cloning is one of the easiest cloning methods in terms of hands-on time, as digestion and ligation can be done in one 30-minute reaction. The destination vector and entry vector(s) are placed in a single tube containing the Type IIS enzyme and ligase. Although the original destination vector + insert may spontaneously religate, this transient construct retains functional Type IIS sites and will be re-digested. In contrast, formation of the desired ligation product is irreversible because this construct does not retain the enzyme recognition sites. As a result, the ligation process is close to 100% efficient. Another strength of Golden Gate cloning is its scalability. Unique 4 base overhangs can be used to assemble multiple fragments - up to 10 fragments are commonly assembled in a single reaction! These overhangs specify the desired order of fragments, and the loss of enzyme recognition sites after ligation favors formation of the construct of interest. Although efficiency may decrease with an increased number of fragments, or the ligation of very small/very large fragments, these problems can be overcome by screening a higher number of potential clones. Golden Gate assembly has a few advantages over other cloning methods. Exonuclease-based methods like Gibson assembly require 20-40 bp of homology at the ends of DNA fragments to specify assembly order, so fragments with 5’ or 3’ sequence homology cannot be assembled using this method, but can be assembled with Golden Gate. The popular Gateway cloning system produces constructs with an attB recombination scar encoding eight amino acids, but Golden Gate assembly can be designed to be scarless. Golden Gate assembly is also less expensive than many commercial cloning methods."

Hello In Kang,

In the past, I had to perform PCR on sequences that form stem-loop secondary structures, like shRNA do. Those sequences have an equilibrium between folded and open structure, and as such they can migrate as doublets. I thus agree that you could sequence both bands to confirm that they are two secondary structures of the same amplicon. Hope it helps.

It is better to look for a vector with these sequences included. You don't want to waste your resources and time cloning 4 inserts into your backbone while also making sure they are in frame with the promoter. I think you can find a vector with the kozac sequence and the secretion tag quite easily. Then you can design your primers for the RBD region with the his tag included and clone the His+RBD insert into that vector.

Hi Sebastián, after digesting with restriction enzymes 5' end will have a P. No worries, it will work.

For those wondering whether it worked, it seemed absolutely fine using the Hanahan protocol! It produced some good competent cells that appeared to have the same stability features as the NEB preparations.

It might be prudent to grow them initially with Streptomycin and Tetracycline to ensure none of the genes that make the 'Stable' phenotype are lost. This also proves useful using these resistance genes anyway, considering usually you wouldn't use drug selection to make competent cells and thus are prone to contaminants. Though further outgrowth I believe won't require this.

Happy cloning!

Not totally familiar with this kit but since most of Maxiprep kits work on the same principle I can try to point out one possible mistake.

After filtrating your Lysate/Precipitate on the syringe you have to add a binding buffer to your filtrate before to apply it on the DNA binding column. Did you do this step ? Omitting it results in very poor binding of the plasmid DNA to the silica column and ends up with super low amount of purified DNA

The problem primarily arises if clones (or phages) have unequal growth rates then you will amplify any bias you have. When you go through the single colony phase although there may be size differences with an overnight growth on plates there is ample opportunity for the slower growers to somewhat catchup with the faster growers. But this is much less true in liquid. But if everything is growing at the same rate then it might not matter so much.

I am having the same issue. Did you problem solve after trying DH5-alpha?

Hi there, With the view of plasmid amplification, one transformant is enough. Are you sure about the amount of plasmid you engage in the transformation? If you get very few clones for one plasmid and loads with the other it might be due to the amount of circular plasmid in the transformation mix. If you want to make sure that the few clones you get are relevant you have to run the control experiment without plasmid to check the behaviour of the recipient strain on selective media.

Assuming your gene is protein coding:

Removing the IVS will probably lower the expression of your protein.

Deleting the IRES could cause major problems, it's there so two different proteins can be translated from a single mRNA. This could result in cells not expressing the selection marker if expression of both your gene of interest and the marker are driven by the same promoter.

Here's some articles:

Hello Chun-Chen Yao ,

Regarding your questions. When you transform the cells with a vector, with a mixture of linear and circular plasmid form, only the circular form will be useful for the bacteria (for the resistance Ab as an example), and the linear form probably will be degraded. The linear plasmid form ligation inside the cytoplasm is unlikely, is more probably that de linear dsDNA will be degraded outside. You have to think that the transformation os linear dsDNA is more difficult in comparison to circular dsDNA (which is more compact).

I don´t know if this question comes from a problem with the E. coli transformation. If it was that the case, you can consult the book: Molecular Cloning: A Laboratory Manual by Joseph Sambrook and David William Russell

If my answer has been useful, please recommend it.

:) :)

And Good luck!

DH5alpha does not require arginine to be added. The argF gene, which is deleted, is a duplicate gene so is not essential. Assuming you have the right concentrations for your chemicals and are following a recipe for a standard medium, then check the glucose concentration you are adding and also the pH. Both of those could be problems. I would also suggest you might try using a different minimal medium just to see if it grows better.

Note that BL21 is RecA+ and DH5a is RecA- so will not grow at the same rate or to the same density as BL21.

As mentioned by Hanna Alalam it may depend a bit on the strain but in theory you should be able to transform nearly any E. coli strain. However whether you get good expression of the GFP will depend upon the construct itself, is there a suitable promoter and appropriate translation signals for GFP expression in E. coli.

Hi,

I think first you should check whether cloning occurs properly. If cloning is confirmed then you will definitely get colonies in antibiotic containing plate.

I haven't personally, but this paper did. Hope it helps!

Hello Rafael,

Have you checked for the basics? Gel concentration, enzymes activity, water quality etc? Sometimes can be simple like that.

Best,

Roberto

AAV2 would be a sensible first option. I've used AAV2 to transduce SY5Y cells before. AAV-DJ would be my other recommendation and often results in higher transduction of a broader range of cell lines.

Interesting question. Following the discussion.

Hi Daniel Dehany Scott ,

at the end, did you find this plamid?

best,

Antonino

Yes, you can do that without problem. In fact you don't even need to heat kill if you are just doing diagnostic cuts. However be sure you are using a buffer system where both enzymes work well.

SpeI should not have been affected by methylation, only Cla (sometimes). If you know the sequence at the two ClaI sites you can predict if it is blocked or not.

Have you thought about verifying your clone by PCR?

Thank you so much Wenyan Xu. Greatly appreciate the advice.

Yes, this sounds normal.

It is totally safe to grow E. coli @ 30°C and 25°C for plasmids preparation. The time to reach culture saturation (stationary phase) will mainly depend on the initial load in your samples and its volume. In rich media supplemented with glucose, you should be able to get enough bacteria in a 12 hrs ON culture, in regular E. coli lab strains.

Are you using a column prep kit for this or just the base solutions?

I agree with your logic but, as with many things, logic can change based on understanding. When I read your statement, it made me think more deeply about the process of plasmid purification.

When you purify the plasmid from a young culture, the amount of DNA is limited in the sample (remember that the kits are designed for a certain number of cells to be processed). This means that the column in this case has a binding capacity that is nowhere near saturation, potentially allowing other salts to bind (think about why DNA binds them - the charge is critical) and be "purified" with your sample. This would explain the arcing that you see when you try and transform the plasmid.

I am not an expert on sequencing but have to believe that the salt contamination would also mess with the analysis.

My suggestion, if you want to experiment, would be to grow a larger volume for the shorter time (maybe several tubes or flasks) and test the preparation from one versus a pooled sample. If the pooled sample has less trouble electroporating, my suggestions are probably correct and you can send it out for sequencing as well.

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Did you solve your problem? pCC1 is a single copy BAC, so you you will get VERY little DNA from a mini prep culture (about 50-100 ng total form 1ml overnight culture).

You can relax the single-copy control of the vector by using a "proprietary" induction solution (arabinose), and get 10-20x more DNA. But you probably chose pCC1 for a reason, is your insert toxic? In that case induction of pCC1 may lead to the very same problem that you hoped to overcome by using pCC1 in ther first place (genetic instability?).

Best if you simply prepare DNA from a larger volume culture. We also had good success, by seeding a lawn of E.coli on chloramphenicol plates, and prepare DNA from the bacteria on the plate.

I do not have an answer but actually face the same question. In my case, the ATG inside the chimeric intron is unfortunately in-frame with my insert, thus there is a contiguous ORF from the chimeric intron's ATG all the way through my desired product. So I'm also wondering if I might look at a potentially altered N-term.

Does somebody have an idea whether the chimeric intron ATG can lead to undesiredly translated product when it gives rise to an ORF (or is even contiguous with the ORF of the insert)?