Bacterial Transformation - Science topic

I was actually thinking of GFP expressing clones. In my experience these are colored in the green/yellow-ish tones... But it can't explain the possible "leaky expression". Have you got PCR positives among the non yellow-ish clones?

Hi Diogo Figueiredo . Thank you for the suggestions. Yes, we will add extra G for the sgRNAs. For the dilution of annealed oligos, I followed from Zhang's protocol. Do you have any supporting documents for your protocol? Thanks in advance!

a replicative plasmid is a plasmid which, once inside its bacterial host cell, will remain in its extra-chromosomal form; its replication is autonomous and does not depend on the replication of the chromosome.

On the other hand, an integrative plasmid is a plasmid which cannot be found in an extrachromosomal form, it must integrate the bacterial chromosome, it cannot replicate autonomously, its replication depends on the replication of the chromosome.

For my understanding, you're describing competent cell preparation, and I'm suggesting that your buffer contained CaCl2 and glycerol that are prepared in Tris buffer, as Tris has its pka as 8.0, by adjusting its pH to 8.0 will provide the best buffer range. While not necessarily in pH 8.0, pH 7.2~8.0 are an acceptable range, pH within this range should not cause any significant damage to cells or DNAs, and the buffer will protect your cells until transformation process is done.

if the stock solution is 1000X then you just calculate volume per volume. For 1 liter of media you would use 1ml of the stock. For 500 ml of media you would use 0.5 ml of stock. So in other words 1/1000 of the volume of media you have.

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.

Yes, you can adapt the protocol for preparing electrocompetent E. coli cells from DH10B to JM109. However, it's important to note that different strains of E. coli may have slightly different requirements for optimal transformation efficiency, so you may need to optimize the protocol for JM109 cells.

Here's a general outline of how you can adapt the protocol for preparing electrocompetent JM109 cells:

By following this adapted protocol, you should be able to prepare electrocompetent JM109 cells for your Gibson Assembly experiments. It's always a good idea to perform optimization experiments to determine the optimal conditions for your specific application.

Hi Dear Teresa Blázquez, Did you try this method? what were the results.

When performing a ligation reaction with one vector and two inserts, there are a few key considerations to increase the likelihood of successful ligation. Here are some recommendations:

1. Vector and Inserts Preparation: Ensure that the vector and inserts have been properly prepared and purified. This includes digesting the vector and inserts with the appropriate restriction enzymes, dephosphorylating the vector (if necessary) to prevent self-ligation, and purifying the DNA fragments from the digestion reaction using a DNA purification kit.

2. Insert Ratio Optimization: Determine the optimal molar ratio of vector to inserts. This ratio can vary depending on the specific experiment, but a standard recommendation is to use a two-fold molar excess of the vector compared to each insert. Adjust the amount of DNA accordingly to achieve the desired ratio.

3. Vector Control: Include a vector-only control in the ligation reaction. This control helps to identify any background self-ligation of the vector or potential contamination of the components.

4. Extended Incubation: Allow for a longer incubation time during ligation. Incubating the ligation reaction overnight at a lower temperature (such as 4°C) can improve the chances of successful ligation by giving the enzymes more time to bind DNA ends and create ligated products.

5. Optimal DNA Ligase Concentration: Determine and optimize the concentration of DNA ligase enzyme in the ligation reaction. It is recommended to follow the manufacturer's instructions or perform preliminary experiments to find the appropriate concentration that promotes efficient ligation without causing excess background ligation.

6. Ligation Enhancers: Consider using ligation enhancers, such as polyethylene glycol (PEG) or dimethyl sulfoxide (DMSO), to increase ligation efficiency. These additives can enhance DNA hybridization and enzyme activity, thus improving the likelihood of successful ligation.

7. Transformation and Selection: Transform the ligation reaction into competent host cells using an appropriate method (e.g., heat shock for bacterial cells). Following transformation, select transformed cells using selective media containing the appropriate antibiotic resistance marker present in the vector.

8. Verify and Validate: Perform colony screening, diagnostic PCR, or DNA sequencing to verify that the desired ligation product has been successfully obtained. This step is crucial to ensure that the correct inserts have been ligated into the vector.

By following these recommendations and optimizing the key parameters, you can increase the chances of obtaining successful ligation with one vector and two inserts. It is always advisable to consult relevant scientific literature or seek guidance from experienced researchers in your specific area of study to further refine your ligation strategy.

upon further reflection, I should add that the answer depends on the nature of the ss DNA. If you have supercoiled plasmid which is denatured by heat, then this is really not going to be single stranded. The two DNA strands will be intertwined even when denatured and upon lowering the heat the DNA will reanneal generating double stranded. If you truly have single stranded DNA then the answers above stand, depending upon the situation.

Are you trying to express your protein of interest? In that case, you might need to use BL21 or some other expression vectors. DH5a in not a protein expression vector, so your pET28a containing the gene of interest might not get expressed, thus everything is growing on the plates!

Hi, for bacmid extraction we proceed as with small plasmid (3ml of preculture/pellet/solution sds NaOH/then acetate de K...phenol/chloroform/EtoH precipitation) exept that we did not vortex (just invert tubes up and down by hand 5-times at each step). However the yield is very small as large plasmids are not in multiple copies in the bacteria (so it is not easy to see them on a gel; they were only used to transfect SF9 cells) If you want to do restriction map for example you will need to start with larger volume. there is kit to extract bacmid (Qiagen, Thermo-fischer....)

Sure you can.

Here is the protocol I use.

Preparation of competent E. Coli cultures

Important: Do not use any antibiotics for DH5a.

Inoculate an LB culture with DH5α cells (directly from the frozen stock without thawing) and grow overnight at 37 °C.

1. Inoculate 5-6 colonies in 100ml SOB medium in 1L flask, grow to OD600~ 0.3-0.6 at 18°C, (250 rpm). Do not exceed 0.6.

2. Incubate on ice for 10 minutes

3. Centrifugate (2,500g 10'min at +4°C).

4. Resuspend the sediment in 32ml of cold TB.

5. Incubate on ice for 10 minutes

6. Centrifugate, 2,500g 10 min at +4 °C.

7. Resuspend the sediment in 8ml of cold TB.

8. Add 560ul DMSO (carefully, slowly, I recommend add it on the tube wall while slowly rotating the tube).

9. Incubate on ice for 10 minutes

10. Dispense the suspension into sterile microcentrifuge tubes (100 ul is enough ph).

11. Freeze in liquid nitrogen

12. Store at -80°C

Transforming a pSOUP plasmid into Escherichia coli (E. coli) DH5α is a common molecular biology technique used for cloning and gene expression studies. pSOUP plasmids are versatile tools that can be used for various genetic manipulations. Here's a step-by-step guide on how to successfully transform a pSOUP plasmid into E. coli DH5α:

i do not know what went wrong with your transformatio, but efficiency is lousy (could be compatents qualit, not allowing 1h recovery, DNA quality,…). Any way, statistics of 3 colonies (in LB+K) vs 0 colonies (in LB+K+I) is meaningless, so I will suggest putting the metabolic burden idea on hold.

Instead, start by picking and plating the 3 LB+K colonies on LB+K+I, see if this solve your imidiate problem (I.e. the colonies are fluorescent with IPTG but less so without).

For the long run, debug your transformation! Take a plasmid that worked well for someone else in transformation and use low amount. Try the instructions here:

It's hard to interpret your gel because there is no ladder, so we don't know the DNA sizes of each band. I would recommend always including a ladder when running a gel.

It looks to me that Trans1 and Trans2 worked as expected because you have a single band. The reason this band is lower on the gel is not from degradation, but because plasmids from bacteria are supercoiled so they migrate quicker through the gel. I'm guessing the Backbone lane in this gel is the linearized backbone that you used in the ligation; since it's linearized it won't be supercoiled, so it will migrate more slowly through the gel.

For Trans3, it looks like you have that same supercoiled plasmid band, but I don't know what the smear above it is.

Well could be the non-structural protein has a temperature dependent interaction with something in your expression vector. NS1 is known to affect lipids while NS4 Modifies the ER.

Your plates look similar to LB without antibiotics as you are getting a smeary sort of lawn growing.

Your Kan is likely expired, and Amp can breakdown quickly. So, you're likely losing selective pressure. That would explain why you aren't getting any plasmid back out after your liquid culture.

Also, if the antibiotics are added when the medium is still hot the antibiotics will break down.

I would suggest making up fresh solutions of your antibiotics, filter sterilizing them, making small aliquots, and storing at -20. Add the antibiotics to liquid cultures right before culture inoculation (don't store liquid LB with antibiotics, it breaks down).

Hope this helps!

Hi Sanjay T D I'm currently trying to clone a 700bp insert into a 18kb vector and I'm having the same issue as you : no colonies. I'm pretty sure the problem comes from the transformation part and not the gibson assembly. I've read a lot about it and apparently 18kb is really big for bacteria if you're doing a heatshock (like I do). people recommand using electroporation instead (if you can).

If you manage to clone your insert please let me know because I'm really struggling. So far I've tried different ratios 1:1, 1:3, 1:10: 1:20 with vector quantities from 100ng up to 300ng.

Have a nice day.

Lauren Woodward Hartzler

Competent Agrobacterium cells:

Transformation:

Skipping Rnase should not impact the transformation. Using 1000 ng is a ton of DNA, you may want to try using a smaller amount. The colonies you are getting look really nice so if the goal is just to have your plasmid in culture then it looks like you succeeded. I agree that the efficiency is quite low. Are your competent cells starting to get a bit old? They generally stay good for about a year at -80.

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.

Daniela Liebsch Thanx for wonderful explaination

It depends on the gene of interest and the type of experiment you are running. Generally speaking, if you need to perform a specialized experiment, it may be necessary to order a multibac plasmid. If you are just performing a basic experiment and your gene of interest is compatible with DH10bacs, then you can go down this route.

I recently sequenced pHIT60 via https://www.plasmidsaurus.com. In case anyone is still looking, Iv'e attached the gbk file!

Hi all,

Thank you for your answers,

I did a restriction digest with enzymes that cut multiple times and indeed, this plasmid has recombined in all sorts of ways except the one I was planning on...

I don't know if any of you have practiced recombineering before, but if you have I would really appreciate your advice regarding how to reduce unwanted recombination events in this type of cloning.

I am using an L-arabinose inducible plasmid for the λRed system. Are NEB10betas good cells for these protocols or maybe Stabl3 would be a better option? Also, would co-electroporating my plasmid at very low concentrations and the linear dsDNA into E. coli (which contains the induced λRed system-plasmid) help in avoiding these undesired recombinations?

Any other thoughts or help on how to avoid this?

Thanks!

Thanks all!

In order to answer your question, some terms need to be better defined. First you need to specify what volume of cells in your transformation. For a typically transformation of 100ul of competent cells is not going to give the same answer as a scaled up transformation with more cells. Also what is your criteria for transformation?

Assuming a normal amount of competent cells, what you would see is a curve if you plot transformants per amount of DNA used starting from the lowest amount tested, until you hit the saturation plateau. The minimum will be 1 molecule, the probability of transformation is low but not zero. Even at sub-ng amounts of DNA you will still get lots of transformants.

Saturation is hit probably in the 10-50 ng of typical cloning plasmid and adding more DNA is likely to only slightly increase number of transformants above this. But this isn't really a maximum, just approaching it. At some point though you may start getting into inhibition with even higher levels of DNA.

Generally cells that have just been transformed need treating very gently and would not be spun down until they have been plated and have formed colonies.16000 is much too high a G force unless spinning cells down to make dna so there is no need for the cells to survive. Why do you need to spin these cells at this stage?

We got WB800N here: https://lifescience-market.com/index.php?main_page=product_info&products_id=64199

Regarding digestion: presence of 250 bp band suggests, that at least part of your plasmid has been cleaved. But this is never 100% effective and at least small fraction of uncleaved vector will markedly affect your result. Most of your clones will be empty.

Dear all,

Thank you for the suggestions, will try them out.

Yes! After the heatshock we incubate the bacteria for 1 hour at 30° (stbl3 strain optimum temperature) in an incubator shaker and then we plate it.

So it is clear that something is not working in your procedure, so you need to do controls and confirmations. The likely things to suspect are:

1) your plates are incorrect (someone added the wrong antibiotic)

2) your plasmid is wrong or there is no plasmid DNA

3) your cells are not competent or are dead

4) you are not following the protocol properly

Most likely the answer is 3, but all should be checked and confirmed until you find the problem.

Recheck your whole process.

Are you saying that a single protein could weigh that much? That would make it about 1.5 million aminos in length requiring an mRNA at about three times that length, not counting the UTRs a both ends... Sounds implausible. Did I misunderstand something you folks were saying?

Simply add low material

Use AMP for bacteria. The KanR cassette is driven by the simian virus 40 promoter meaning it is for eukaryotic expression. The KanR gene is used for G418 selection.

Zymo purification kit is a great option for you. Just make sure that your product is completely digested. I recommend overnight digestion just to be 100% sure. After the overnight digestion uses a purification kit to cleanup your linear product. It is not necessary to use a gel extraction kit as you may lose some of your DNA during the process. Keep in mind that for Pichia transformation you will need at least 1ug DNA for a successful transformation.

What sort of contamination are you seeing? Is it fungal or bacterial?

I would suggest getting all fresh medium (liquid culture, plates) and seeing what happens if you do a mock transformation (to see if your cells were contaminated to start with).

Do you use a frozen stock of competent cells or make them fresh? One time I had to re-make competent E. coli stocks because ours were contaminated with what looked like Staphlococcus (the person who made the cell stocks had a really bad cold)

Congrats! So glad I could help! :)

Dear All,

The problem is fixed; it's caused by plasmid aggregation I guess

because when I try to linearize, it somehow shows a single band with the correct size in the gel.

Sailee Asolkar You'll still use erythromycin selection like you've been doing. Even in the event of a successful transformation, the overwhelming majority of your cells will remain untransformed and erythromycin sensitive, and will adopt that morphology. Basically I am saying the cell morphology is irrelevant to your failed transformation problem and not to worry about it.

Are you getting uniform colonies after transformation? I had a cell clumping issue one time and it turned out the person who made the competent cells contaminated them with Staph aureus (they had a nasty cold when prepping the cells).

I usually use room temp SOC, that part should be fine.

Another possibility for the clumping is your shaker is set to a lower rate, allowing cells to settle.

Old cells that are less viable would only contribute a tiny amount of dead cells to the vial, that doesn't seem a likely source for clumps.

Mitchell Pallett Katie A S Burnette thank you for your answers and recommendations

Thank you, everyone for your responses!

Nikolaos Mastrodimos

You are welcome, and let me know your new results of using another Gateway destination vector, to see if it works. Good to talk to you, and good luck.

If you take care of avoiding microbial contamination, they can be kept forever, preferably in the refrigerator, but this is because they need to be cold when you want to use them to prepare competent cells.

Avoid autoclaving glycerol and sugars together with divalent cations (I am not sure about glycerol, but certainly for sugars, which may form toxic byproducts like furfural).

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.

Hi Ahmet Can ,

2 possibilities are that the Amp is getting old and selection is not strong or that the insert is toxic to cells ( try incubating at 25c may help).

The troubleshooting guide from NEB at

suggests some good control samples to run to assist troubleshooting

may be of interest.

Hello Hasan Nasrullah, Ana Crnkovic, Chankit Giri,

thank you so much for your valuable recommendations.

actually, this is the first time to do transformation. I followed the same steps mentioned in this link:

https://www.addgene.org/protocols/bacterial-transformation/

the plasmid I used is the PEX-1 vector bearing Ampicillin marker and for bacterial recovery, I used the LB medium, not SOC.

so, as you mentioned, I need to grow DH5A in LB without antibiotics to make sure of its quality.

Joseph Ayariga I am using pT7CFE1-CHis to generate a fusion protein, which we will detect by using an anti C-His antibody.

The insert have a size of 2796bp and the vector has 3627bp.

Interesting

For standard transformation purposes your cells should be fine. If you try again with the 1 hour outgrow time that should lead to a higher efficiency as it's about 1.5 more doublings than 30 minutes. I really haven't noticed a difference between using SOC and LB, both seem to work well.

No, you did it right. It's one of the things they don't teach in classes and don't write into papers: When you try to express any random protein from a higher kingdom in E. coli, most of the time, it does not work. This appears to be due to sequence / structure issues that have evolved somewhere in the Eukaryotic lineage, that E. coli has a very hard time dealing with.

So, you found one of those. Congratulations. Now what? There is a long tail of experiments you can do, to "make it work". If you go through all of that, you'll have a somewhat better chance to succeed than fail, but not much better than 50%. Things to look at:

Can you find anyone else who has expressed it -- what did they do? Make a fusion protein with an N-terminal leader, such as MBP. Make the gene synthetically and "optimize" codons for E. coli. Think about natural stabilizers of your protein. Is there a co-factor you can provide? A binding protein you can co-express? Finally, have you added a protease inhibitor cocktail? (It's rare to have proteolytic degradation in BL21, but it can happen. More likely you make the his-tag and then the ribosome derails because it doesn't like something about your protein...)

If none of these do it, and you still want your protein... You'll have to move up the chain into eukaryotic expression.

It’s 2021 And we have started using this cloning kit. Initially we had problems like multiple bands appearing in the cloned products. However this was solved with increasing the ligation time. Now the problem is that, it works completely fine with smaller sized inserts (below 1kb) but not with larger ones. We are still getting multiple bands.

can any one suggest me?

Mercedes Vazquez Can I as you what plasmid you are talking about? Is it pCP20?

Many thanks?

I think you're having a few different issues here.

Firstly, you should probably run more DNA in each lane to better visualize the bands. For example, your uncut plasmids should produce 2 distinct bands (circular and supercoiled).

Secondly, the size of the extracted genes does not match the expected size. pGEM-T Easy is a 3kb plasmid and your undigested bands run at >20kb, implying a 17kb insert. However, the bands in the digested lanes are all <1.5kb, far smaller than expected.

I would strongly suggest that you verify the plasmids you are using by digest screening or sequencing.

Those almost look a 100% like satellite colonies, which indicates that you are incubating for too long. Especially if you are working with ampicillin. Your agar cracking indicates to me that you are incubating for too long as well. The satellite colonies are bacteria that appear ampicillin resistant, but due to the overincubation, the ampicillin in the area around your true colonies is degrading, which gives non-ampicillin resistant colonies the chance to grow.

M13 isn't a plasmid replication origin of replication, it's for packaging into M13 phages which used to be common. The actual origin of replication on that plasmid is probably elsewhere.

There's a bunch of reasons why there could be a difference in transformation efficiency, and unfortunately you might not get a straight answer without more information and/or doing a bunch of experiments that I assume aren't your primary focus.

Some that I thought of:

Hello Rohit,

I have recently did the experiment where i use PCR product as a template. In this situation i had good product when you dilute the template (PCR product) to 1:50 or more. In theory it should be more diluted sometime to more then 1:100 or more. I can suggest you to have some parallel with dilution start from 1:50 and higher for template in your PCR reaction. I read that if you use undiluted or less diluted template (PCR product) for reaction, in sometime inhibit the PCR reaction because of high concentration of template. So you can try to work with some higher dilutions. Good luck.

Best,

Mradul

The frequency with which markers separate, that is, (1 − CTF), is a measure of distance between markers. For example, if two markers show 20% linkage, they might each transform 1% of the cells in a population, but 20% of the cells transformed for one marker (0.2% of total cells) will also carry the other marker. If the markers are not closely linked, they will be separated by the dispersive processes and enter cells separately and randomly. For an individual marker transformation frequency of 1%, the CTF would be 0.01% of total cells (or 1% of cells transformed for one marker would be transformed for both markers). CTF refers to co-transformation frequency. It is a useful index of transformation efficiency.

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.

do your pcr primers to check uptake go from vector into insert or sre they just insert primers?

If just insert is it possible that you have spread template dna on the selection plates with the cells and your positives are just wetting effect on the cells so initially pcr positive but when grown up and diluted the colonies are negative because they always were

I do a 1 hour shake in SOC as a "recovery" immediately after the transformation & before plating on selective medium. SOC is a high-sugar growth broth. It typically comes in most transformation kits.

Yuan-Yeu Yau <_{Were you talking about your research plant species, Passion Fruit (know this from your other thread)? Their genome has not sequenced? >}

Yes; about Passion fruit,

As the genomes on passion fruit published this year but started work on passion fruit 2 years ago by RNA-Seq. At that time no genome reference, so selected the target genes (CDS), for example, XYZ on the basis of Arabidopsis and RNA-Seq, and constructed the binary vector with target genes.

When compared those target genes (CDS) with genome and found that some gene has more than 90 % identity RNA-Seq and Genome, but some genes < 40%, so will its influences while publishing?

Or should re-assemble the RNA-Seq with the passion fruit genome? (as before RNA-Seq has higher similarity with Populus Trichocarpa")

Hello, as DNA is a highly hydrophilic molecule, normally it cannot pass through the cell membrane of bacteria. Hence, in order to make bacteria capable of internalizing the genetic material, they must be made competent to take up the DNA. This can be achieved by making small holes in bacterial cells by suspending them in a solution containing a high concentration of calcium. Extra-chromosomal DNA will be forced to enter the cell by incubating the competent cells and the DNA together on ice followed by a brief heat shock that causes the bacteria to take up the DNA. Additionally, a poorly performed procedure may lead to not enough competence cells to take up DNA. The divalent cations generate coordination complexes with the negatively charged DNA molecules and LPS, the monovalent that you mentioned can't. DNA, being a larger molecule, cannot itself cross the cell membrane to enter into the cytosol. The heat shock step strongly depolarizes the cell membrane of CaCl2-treated cells. Thus, the decrease in membrane potential lowers the negativity of the cell’s inside potential which ultimately allows the movement of negatively charged DNA into the cell’s interior.

Hope this info will help you.

Goof luck!

Hello, first did you get any colony using your conditions?

if not you can try 1:5 (vector: insert) molar ratio using 200 to 500ng total amount of backbone. Then you can transform up to 5uL in your 50uL aliquot (should not exceed 10% of volume of cells).

Hope this helps.

You want to check your construct to ensure you don't have any toxic gene like ccdB. If you do, using DH5alpha is not ideal. You will want to try competent cells like DB3.1 or ccdB survival competent cell.

If even the control plasmid is not transforming well then the problem is most likely either the competent cells or the plasmid DNA. Do the appropriate controls with a different plasmid (but same antibiotic resistance) with your mach-1 competent cells, and your plasmid DNA into some different competent cells to see which is not working well.

Your protocol is fine. I presume you have confirmed that the plasmid you are using is actually Amp resistant?

I have a very similar problem, rescuing a (bigger) plasmid of a maxiprep, hope someone can help us