Molecular Cloning - Science method

I have been using NEB Hifi Gibson assembly for a couple years now and I've been quite happy with it. I regularly make plasmid constructs with 4-8 fragments, and always >1/4 of the colonies are "perfect," while the remaining ones may have some SNPs at the joining sites, or be misassembled due to a repetitive region.

Some colleagues said that Golden Gate has even higher efficiency. That even with 8 fragments, it is normal for 50-100% of colonies to be perfect. Is that true? Is Golden Gate that perfect?

I want to insert a gene in a vector using restriction cloning, but the enzyme that I have to use has three restriction sites in the vector. It is imperative that I use only this enzyme and no other, so I can't use a different restriction site or other enzyme. Can someone help with this issue?

I have tried partial restriction digestion with different units of enzyme as well as different time periods of incubation but haven't got a single band. The enzyme is cutting at all three sites whatever conditions I try in the partial restriction digestion.

My PI wants to use expression plasmid for E. coli with two cloning sites. We have pColA-Duet-1 and pACYC-Duet, but in those are both of the sites inducible by IPTG (if I understood it correctly). Are there any plasmids that would have two different inducible promoters?

I'm trying to clone some large pieces of cyanobactrial gDNA (sometimes even more of them) into a plasmid via fusion PCR. Then, I transform it into E. coli TOP10, as my PI says it works better (for HiFi DNA assembly).

However, I quite high percentage of my positive clones had some problems in the plasmid elsewhere (like missing I think 2.3 kbp of the plasmid; or some mess in another case etc.).

It is true, that when I compared TOP10 and DH5alpha, TOP10 had many more colonies (more than 10-times). However, I think that DH5alpha should be less prone to DNA alterations, right? So should I use rather DH5alpha and hope that I will get at least few colonies, but they will be positive?

Hello there

I have some difficulties in my cloning project, and I'd appreciate using your experience in this field.

I want to clone my insert (2.8 kb, cloned in pUC57) into a manually engineered plasmid for CRISPR (9 kb with pUC plasmids backbone) in XhoI restriction site.

For the experiment, I received my synthetic insert in pUC57, then, I prepared my insert with PCR (with speedy pfu as a polymerase with proofreading). (Notice: I can't prepare my insert with enzyme digestion and purification because my digested insert and linear plasmid have same size)

Additionally, I extracted my plasmid by Qiagen kit and eluted with nuclease free H2O. I used Xho1 treatment for both Insert (PCR product) and Plasmid in my digestion process, and I tried the gel purification for preparation of my XhoI digested-insert and eluted with nuclease free H2O. In order to do single digestion cloning, I needed to use alkaline phosphatase treatments (Roche) after plasmid precipitation and deactivate it in different ways for multiple cloning set up. So, the XhoI digested plasmid was treated with Alkaline phosphatase and deactivated in different ways including incubation at 65°C, gel purification, and clean up in independent cloning experiments.

For the ligation step, I tried 48h/4°C and 24h/16°C incubation and pre-warming of Insert+ plasmid in 45°C and 65°C, ice incubation, and then adding ligation buffer and ligase (Takara and Roche ligase were tested). I checked my ligase and it works well in different project.

Recently I tried my cloning process with double digestion by forward primer changing (replacing NcoI instead of XhoI) in preparing Insert by PCR, but I couldn't get any cloned plasmid.

I have tried different protocols, and I welcome your comments and experiences in this regard.

In many cloning protocols, researchers introduce an intermediate cloning step using a vector like pGEM-T before transferring the insert into the final destination vector. Could you elaborate on the rationale behind this approach and the specific advantages it offers, especially when considering the additional time and effort involved?

What are the specific advantages of this approach?

Thanks

My vector size is 5.1kb and insert size is 4kb. Initially, I performed digestion of the insert from a different plasmid and the vector using KpnI-NotI. I checked them on the gel, excise and extract the band from the gel. I aimed ligation 1:3 and 1:5 at 4C and 16C overnight and RT 1hr ligation. I used fresh enzyme and buffer. I performed transformation in Stellar cells. However, I got very few transformed colonies on plates where ligation was performed 1:3 at 16C ON. Recovery of Stellar cells was great. Unfortunately, the transformed colonies do not contain the insert. They carried the empty vectors. Any thoughts?

I know, IRES enables the coordinated co-expression of two genes with the same vector, used for the expression of two proteins separately.

But I found two kinds of IRES sequences in my plasmid database and literature. Here it is:

IRES:

TCCCTCCCCCCCCCCTAACGTTACTGGCCGAAGCCGCTTGGAATAAGGCCGGTGTGCGTTTGTCTATATGTTATTTTCCACCATATTGCCGTCTTTTGGCAATGTGAGGGCCCGGAAACCTGGCCCTGTCTTCTTGACGAGCATTCCTAGGGGTCTTTCCCCTCTCGCCAAAGGAATGCAAGGTCTGTTGAATGTCGTGAAGGAAGCAGTTCCTCTGGAAGCTTCTTGAAGACAAACAACGTCTGTAGCGACCCTTTGCAGGCAGCGGAACCCCCCACCTGGCGACAGGTGCCTCTGCGGCCAAAAGCCACGTGTATAAGATACACCTGCAAAGGCGGCACAACCCCAGTGCCACGTTGTGAGTTGGATAGTTGTGGAAAGAGTCAAATGGCTCTCCTCAAGCGTATTCAACAAGGGGCTGAAGGATGCCCAGAAGGTACCCCATTGTATGGGATCTGATCTGGGGCCTCGGTGCACATGCTTTACATGTGTTTAGTCGAGGTTAAAAAACGTCTAGGCCCCCCGAACCACGGGGACGTGGTTTTCCTTTGAAAAACACGATGATAA

IRES2:

CCCCTCTCCCTCCCCCCCCCCTAACGTTACTGGCCGAAGCCGCTTGGAATAAGGCCGGTGTGCGTTTGTCTATATGTTATTTTCCACCATATTGCCGTCTTTTGGCAATGTGAGGGCCCGGAAACCTGGCCCTGTCTTCTTGACGAGCATTCCTAGGGGTCTTTCCCCTCTCGCCAAAGGAATGCAAGGTCTGTTGAATGTCGTGAAGGAAGCAGTTCCTCTGGAAGCTTCTTGAAGACAAACAACGTCTGTAGCGACCCTTTGCAGGCAGCGGAACCCCCCACCTGGCGACAGGTGCCTCTGCGGCCAAAAGCCACGTGTATAAGATACACCTGCAAAGGCGGCACAACCCCAGTGCCACGTTGTGAGTTGGATAGTTGTGGAAAGAGTCAAATGGCTCTCCTCAAGCGTATTCAACAAGGGGCTGAAGGATGCCCAGAAGGTACCCCATTGTATGGGATCTGATCTGGGGCCTCGGTACACATGCTTTACATGTGTTTAGTCGAGGTTAAAAAAACGTCTAGGCCCCCCGAACCACGGGGACGTGGTTTTCCTTTGAAAAACACGATGATAATATGGCCACAACC

Somehow, I want to know what is the difference on the expression level of these two sequences. Someone said IRES2 will decrease the expression of the second gene compared with IRES, is it true? Could IRES keep same expression level of two genes (I know people will suggest 2A peptide, but I do not want to introduce any amino acids on my protein)?

I am cloning a small fragment into my vector harboring Cas9 gene using BplI enzyme for restriction digest. I linearized my vector and treated with rSAP before performing gel extraction to excise vector fragment. (The vector size is 11.8 kb.) My fragment is 20 nucleotides in size and is duplexed. After restriction digest, the vector concentration is generally low less than 30 ng/µl due the enzyme inability to completely cleave DNA.

After attempting multiple ligations and transformations, it appears that there is approximately an equal amount of colonies on both negative control plate and experimental plate (vector + insert). When checking to see if I had gotten any positive transformants using PCR as a diagnostic tool, it resulted in no band at expected size.

I have altered the ligation mix using 1:3 vector to insert ratio. Used different ligases (T4 DNA Ligase and Instant Sticky Ends Ligase) and different transformation protocols.

My question is, why is there an abundance of colonies on my negative control plate and why am I struggling to obtain successful transformants? Could it be that my vector is the problem or is it my duplexed oligos? I tried transforming cells in XL-1 Blue competent cells and Beta-10 competent cells and had gotten the same results. Where am I going wrong?

Ligation Mix:

vector 3 µl (81.6 ng) + insert 5 µl (250 ng) + T4 Ligase Buffer 1 µl + T4 DNA Ligase 1 µl.

For the negative control ligation mix:

vector 3 µl (81.6 ng) + ddH2O 5 µl + T4 DNA Buffer Ligase 1 µl + T4 DNA Ligase 1 µl.

I incubated the ligation at RT for 1 hour.

In the gel image: Lane 1: 1 kb DNA ladder; Lane 2: Negative Control; Lane 3-10: Colonies from vector + insert plate; Lane 11: 1 kb DNA ladder. Gel is 1% TAE.

I am trying to clone ~3.8kb insert into a ~4.7kb plasmid vector. Things are fine upto transformation. However, during screening when I am isolating the plasmid, I am getting only one band(sometimes 2). Also this single band is larger than the clone is supposed to be. Further, when I am doing diagnostic digestion with the plasmid, I am getting a single band after double digestion. Can anyone help me with the troubleshoot please?

Hi,

1. Does any one know what is the maximum length of amplicons we can amplify using PCR? I need to amplify fragments for assembling a big vector (approximately 100kb).

2. Do you think it's achievable if I amplify 10 genes of approximately 10kB and stitch them together using GIBSON assembly?

Any recommendations/suggestions are appreciated.

Thanks

We have constructed several plasmids that express HA- tagged proteins. At first, all the protein expression of these plasmids were confirmed by WB. But now, after several rounds of amplification of these plasmids, the expression of HA tagged proteins could not be detected. Does anyone have this problem?

I made some competent cells, but using those cells for molecular cloning will produce a lot of wrong plasmids, and the colonies grow very few, so will competent cells affect molecular cloning to construct plasmids?

Hi Community,

I am planning to express 6 human proteins in a mouse cell line. They are from 400 to 800 bp long. I want to use three vectors, each containing two CDS connecting with 2A peptides, followed by IRES promoter controlling various fluorescent proteins. However, I am just wondering if having three of them would affect the efficiency of the processing of 2A peptides and causes a severe downregulation of these genes. Transducing 6 genes separately means I have to go through many more clones to get one that expresses all due to lack of selection.

Has anyone have any experience on multiple multicistronic gene expression?

Could anyone please explain how plasmid incompatibility mechanism work in transformation in molecular cloning?

I would like to clone a large (~40 kb) plasmid.

I am wondering if I should use a high copy or low copy backbone for the plasmid?

I would much prefer high copy number to get greater DNA yield and purity, but I have heard low copy greatly improves stability of large plasmids.

Is using low copy number important for large plasmids?

I'm seeing a lot of unexpected errors in one of my PCR products and I don't know why.

I've been using Q5 polymerase for all my cloning for years now and it's always been great -- I've never seen errors in my PCR products. However, recently I tried to clone something. I got many colonies. I picked some colonies and sequenced them. To my surprise, one of the DNA fragments is always filled with errors. I don't know why. For 15 out of 16 clones, there were 2-5 mutations per clone.

The mutations are real and not an artefact from poor sequencing. The sequencing was very clean. And I sequenced by Sanger sequencing and nanopore sequencing: same sequencing results.

More details for the fragment with errors:

I am sure my reference sequence is correct

I never saw the same mutation twice -- all mutations are different.

I did 25 cycles

The product is ~3 kb

The PCR product is actually the plasmid backbone (I had to change a part of my plasmid backbone)

I did other PCR reactions at the same time and those had no mutations

I am seeing 2-5 mutations per clone.

The template was an old digestion of my vector that I've previously cloned with, with no errors.

Roughly half the errors are g>t, half are c>a, although I did see one g>a

What could cause such a high error rate?

Hello everyone!

I am trying to stablish cell lines stably overexpressing PIWIL4 protein for my research project.

For this purpose we've purchased pLJM1-EGFP (https://www.addgene.org/19319/) plasmids in which PIWIL4 ORF was inserted into NheI cloning site (plasmid map attached). These plasmids contain EGFP as tag/fusion protein, thus EGFP fluorescence is expected in transfected cells. We are using HEK293T cells as packaging cells to produce lentiviral vectors, transfecting them with either empty (pLJM1-EGFP) plasmids or plasmids containing our insert along with structural plasmids for lentiviral production (pMD2.G and psPAX2) using Lipofectamine 2000.

When evaluating EGFP fluorescence in HEK293T 24 and 48h after transfection, only cells transfected with empty plasmid are positive. The experiment was repeated twice increasing the concentration of plasmids, and the same result was observed. Also, when transducing the target cell line using supernatant from HEK293T cells transfected with empty vector, they efficiently showed EGFP fluorescence, showing that the system and the protocol are working fine. Plasmid sequence is attached as well as a figure showing a diagnostic digestion with EcoRI and AgeI, showing that the plasmid sequence seems OK.

Does anyone have any clue to why EGFP fluorescence is not observed in cells transfected with PIWIL4-containing plasmids? We will try to perform western-blotting to check the expression of PIWIL4 on these cells and see if the problem is with expression of EGFP, and not the insert. However, I was wondering if PIWIL4 might inhibit CMV promoter and its own expression in these cells, since PIWIL4 is known to target and epigenetically repress LTRs in endogenous retroviral sequences. Is it plausible? If so, does anyone have references to corroborate that?

Thanks in advance!

Hello all

So I am trying to express a gene in a mRuby2 containing Destination vector (pCAGGS nDEST mRuby2). My question here is, does the attb (the resultant of LR reaction in destination vector) affected the final expressed product, given mRuby2 is at the c-terminus of the expressed gene.

I can introduce a Gly-Ser linker between gene and mRuby but I was wondering if attb affected the structure somehow.

Thank you and I really appreciate your inputs on the same.

Hello everyone,

I am creating transcriptional reporters by cloning a promotor to a modified GFP in a plasmid. The GFP is modified by the addition of 13 amino acids at the end of it to render it less stable (if interest, see paper DOI: 10.1128/AEM.64.6.2240-2246.1998 ). I get the plasmid with the insert (promotor + GFP), but, there are always nucleotides' substitution in the 13 amino-acids tail added to the GFP. The mutations vary within the clones sent for sequencing. I have send the PCR product of the modified GFP used before the cloning, it does not have any mutation.

Where do the nucleotides' substitutions come from ? Any idea ?

Hi all, just a small question.

I run DNA gel using NEB 6x DNA loading dye as the loading dye and Gel red as DNA stain. Sometimes when I have very heavy DNA bands (e.g., after PCR) I can even see the red DNA bands (a bit faint though) directly in the agarose gel without using UV light. What could cause this? As far as I know, the loading dye serves to track the electrophoresis progress and precipitate the DNA sample into wells but has no ability to bind to DNA, while the DNA stain does help visualise DNA but is only visible under UV (at least Gel red requires it). So I am a bit puzzled here.

A lot of answers/literature online say that T4 ligation of >3 fragments is very difficult/occurs at low efficiency.

However, I also see reports/protocols everywhere of Golden Gate being used to assemble dozens and dozens of fragments at high efficiency. There doesn't seem to be that much difference between standard T4 Ligation and Golden Gate except that Golden Gate does the restriction and ligation in one step.

Why is Golden Gate so efficient for 6-24 fragments, but standard T4 ligation is difficult with that many fragments?

My own experiences also suggest Golden Gate is better, but I don't know why:

I tried assembling 4 fragments with standard ligation (after digestion) and got no colonies after several tries, or sometimes some colonies, but the same number of colonies on the "no insert" control plate.

I tried assembling the same 4 fragments with Golden Gate and got ~500 colonies, and only 1 colony on the "no insert" control.

What's the minimal U6 promoter sequence to combine with the multiplexed TetOs for the optimal tTA mediated transcriptional activation (which means the least U6 basal leak without the presence of tTA) ?

Dear All,

I have a particular gene of interest from a wide range of mammalian species that I have inserted into plasmids. I want to check how this gene affect a particular mechanism. However, our lab don't have the cell lines from most of these species, e.g., elephant, bat, cetaceans etc.

[1] Can I use hek293 or BHK-21 cell lines, for example, to study a Bat-gene or elephant gene?

[2] If the first is possible, what happens to the endogenous gene in HEK293 after I transfect with the exogenous gene of other species? How does vector carrying Bat-NRF2 transfected into HEK293 affect the expression of endogenous NRF2 in HEK after transfection?

[3] I am yet to see it in publications so I am not sure if it is defensible.

I will appreciate your advice and suggestions,

Thank you.

I am planning on ordering a number of DNA sequences that range in size from about 50-200bp, to be cloned into a vector. I think the best/ most economical way is to order the sequences as ssDNA, that are flanked by a pair of 'universal' primer binding sequences, so that I can amplify all of them with a single primer pair. I could use a random pair of primers that have used previously, but I was wondering if there is a more commonly used pair of primers for this purpose?

The alternative is ordering it as two ssDNA sequences and annealing them, however this would be considerably more expensive. It would also give me a fairly finite amount of DNA.

Below is a workflow schematic for what I am referring to.

primersite1 - RE site - DNA sequence - RE site - primersite2

primersite1 - RE - DNA sequence - RE - primersite2

primersite1 - RE - DNA sequence - RE - primersite2

- RE - DNA sequence - RE -

- RE - DNA sequence - RE -

I have been using lipofectamine (lipo 3000) for transfection of mammalian cells, and now considering whether I should try calcium phosphate-mediated transfection, to obtain better transfection efficiency. Does anyone has exeperience comparing these two methods? Thanks.

Hi All,

Ive been constructing some plasmids with 4-12 pieces at a time pretty regularly using golden gate assembly for about 6 months now, but its getting to be a bit cumbersome due to relatively low efficiency of some of the constructs and really time consuming construct planning and trouble shooting. Someone at my work mentioned that they've had wild success with cloning difficult constructs (over 20 constructs, ~30kbp, 8-12 piece assemblies at >30-80% correct sequence) in yeast where all other cloning fails. It sounds almost too good to be true. The planning is easier, the reactions are simply TAR reactions, the only downsides i see are (1) the need to transform back into bacteria for large scale DNA preps and (2) long incubation timelines after transformation.

I was wondering whether anyone prefers doing all their cloning in yeast transfer vectors and then transforms back into bacteria? Am I naive in assuming this could save me massive amounts of time in construct planning and cloning execution?

I have similar colony numbers for my self-ligation (EcoRV + Cla1) and my re-ligation (EcoRv+BamH1+Cla1 with two inserts).

My No-ligation plate has no colonies at all.

I wonder whats wrong with my cloning.

I did Digestion at 37 degree

I did ligation at 16 degree overnight with ratio vector 1: inert 3:inert 3.

I did vector de-phosphorylation with SAP for 30 mins.

I did transformation using DH5α competent cells

First time I did digestion for 2 hours + SAP enzyme 0.5 in 50ul and I repeated the same protocol with digestion overnight + SAP enzyme 1ul.

I want to synchronize mouse embryonic stem cells in G2/M phase in order to study transcription of a gene of interest during mitosis and throughout the cell cycle but I'm a having a hard time finding a protocol for this.

Dear fellow scientists!!!

I have been doing cloning lately and have encountered some problems during my experiments, for example, I can get none or very few colonies after transformation into competent cells. Another issue is I inspected low Renilla values in Dual-luciferase assay. Hence, I decided to open a discussion here in order to get some useful advice from all of you.

Let me briefly explain my cloning protocol, and if you found any errors or want to recommend a better technique, please feel free to let me know.

1. gradient PCR.

5uL of 5X buffer

2uL of 2.5mM dNTP

1uL of 10uM of primer of interest

16.5uL of 2ng/uL genomic DNA

0.5 uL of Prime STAR GXL Polymerase

--> The total volume is 25uL (for 1 reaction)

--> Running condition: Activation (98*C for 1min), Denaturation (98*C for 10sec), Annealing ( 50*C - 54*C - 58*C - 62*C for 15sec and 40 cycles), Extension (68*C for 1-3 mins _ Depending on the size of primer: 1KB primer - 1 min or 2KB primer - 2 min), Termination (68*C for 3 min), and cooling down at 4*C .

--> Then running the gradient PCR products in 1% gel electrophoresis to check the quality of primers and the best annealing temperature.

2. Double Digestion (DD):

A. For PsiCHECK-2 :

5uL of 10X CutSmart buffer

1 ug of uncut vector

2uL of Restriction enzyme 1 (XhoI)

2uL of Restriction enzyme 2 (NotI-HF)

Then make up the total volume to 50 uL with pure nuclease-free water

--> Incubate the mixture for 3 Hrs. Then add 1 uL (10U/uL) of alkaline phosphatase CIP and incubate for another 1 Hr.

B. For PCR products:

5uL of 10X CutSmart buffer

41uL of PCR products

2uL of Restriction enzyme 1 (XhoI)

2uL of Restriction enzyme 2 (NotI-HF)

--> Incubate the mixture for 3-4 Hrs.

--> Run both PsiCHECK-2 and PCR products in 1% Gel electrophoresis

--> Then cut D.D band under UV light and extract DNA with DNA Gel Extraction S&V kit

3. Ligation

(The insert: vector ratio is 3:1 or 5:1)

2uL of 10X buffer

2uL of cut (DD) Psicheck-2 vector

(X) uL of DD insert [ X value is according to the insert: vector ratio]

2uL of (200U/uL) T4 DNA ligase

Then make up the total volume to 20uL with pure nuclease-free water.

--> Incubate at 16*C for 1-2 days. [ Note: I incubate at 16*C for a whole day in a 96well thermal cycler machine and then I take it out and store it in a 4*C refrigerator before going home. Then in the next day, I incubate it again at 16*C for a whole day.

4. Transformation:

(I use DH5-alpha as a competent cell bought from a company that recommends that this competent cell is a non-heat shock transformation cell, but is required to heat shock the cell for a vector larger than 6KB)

50uL of competent cells + 2.5uL of ligation mixture

Ice incubation (20 mins)

Heat shock at 42*C (30 - 40 sec)

Ice incubation (20 mins)

Add 450uL of LB broth to recover the competent cells after heat shock

Incubate in a shaking incubator (37*C, 1H)

Spread 200uL of the above mixture into an agar plate

--> Incubate agar plate overnight.

*** However, I got none or very few colonies on the next day!!!

Questions:

1. Is it possible that the double digested products are mutated or damaged because of the direct exposure of UV light when I cut the band ??

2. Is the amount of DNA is too much or too low in the ligation process?

3. Could you kindly recommend to me some advice related to the cloning process that works well in your laboratory??

I am looking forward to hearing from you!!

Your advice would be highly appreciated and helpful in my study!!

Thanks in advance!

I have a low copy plasmid (ori=pBR322) and I wanted change it to high copy (like ori=pUC).

This paper seemed to say that only one mutation was required to increase the copy number:

However, I made this mutation in my plasmid and it didn't change the copy number. My plasmid does not appear to have rop either.

What are the key mutations between pBR322 and pUC that cause pUC to have such a high copy number?

Hello everyone,

It is as the title says. To describe my experiment in more detail, basically I want to compare the changes in transcriptomes between the three conditions (all in organoids):

A) double KO of gene 1 and gene 2,

B) single KO of gene 1,

C) single KO of gene 2

(+ necesary controls)

I am doing the KO's with LentiCrisprV2, and will be doing single cell sequencing to compare transcriptomes. However, in order to decrease costs for single cell sequencing, I want to put all three conditions together. Like so:

1) seed organoids

2) infection with two viruses: one to KO gene 1, the other to KO gene 2

3) results in a bunch of organoids where some are double KO, some are KO only for gene 1, and some are KO only for gene 2.

In order to differentiate between cells that are KO for gene 1 vs. cells that are KO for gene 2, I need to differentiate the two LentiCrisprV2 plasmids somehow. since the sequencing is based off mRNA, the mRNA transcripts need to be different. We want to do 5' sequencing (3' sequencing has a bunch of other project-specific limitations that are too long to get into), meaning whatever sequence that differentiates the two plasmids should be on the 5' end.

Therefore, I'd like to be able to add a FLAG tag, His tag, or other similar tag to the 5' end of the Cas9 sequence of one of the LentiCrisprV2 plasmids. Is anyone aware of whether this affects the performance of the Cas9, and how best the tag could be cloned into the plasmid?

Thank you very much for your help!

Hello !

How can I eliminate primer dimers accumulation in pcr?

Thanks in advance.

Hi everyone,

I'm try to clone a gene into a lentiviral plasmid by pcr + digestion + ligation.

After the ligation with NEB Electroligase, the bacteria (ElectroMAX™ DH10B Cells) are electroporated with the ligated product and plated on Ampicillin-containing LB Agar plates and grown overnight.

The colonies are then inoculated into 5ml LB and, after another overnight growth, miniprepped to screen the plasmid by digestion.

The issue is that, very often, the plasmid isolated from the colonies has nothing to do with my plasmid. In particular I expect a band around 12Kb (see first lane on the gel) while the isolated plasmid is around 2-3Kb (all the other lanes on the gel).

How can these bacteria survive to ampicillin both on the plate and in the LB?

What is this "contaminating" plasmid?

How can I get rid of these colonies?

Many thanks to everyone who will try to help!

Normally, ecoli miniprep cultures are grown for around 12-18 hours at 37C.

However, due to my schedule, I won't be able to harvest the cultures for >24 hours. I think >24 hours is too long to grow ecoli cultures at 37C. I cannot freeze the cells and miniprep them later -- I will not be in lab at all for >24 hrs.

I want to grow them shaking at lower temperatures to slow their growth so I can harvest them later. However, I do not know what growth duration is appropriate at these temperatures.

How long should ecoli be grown at 30C and 25C to get something similar to 12-18 hrs at 37C?

I transformed a bacteria with an integrating plasmid. I grew up colonies in broth and then did colony pcr to genotype them.

I did two pcrs:

1st pcr: Should only give 1 kb band from transformed bacteria

2nd pcr: Should only give 1 kb band from wild-type bacteria

Unfortunately, for all my cultured colonies, both pcrs showed 1 kb bands. This indicates my cultures are a mixture of wt and transformed.

I was very careful in picking the colonies and I do think the cultures are truly a mixture because I saw both bands even after passaging the bacteria a few times.

The only solution I can think of is streaking out the culture for single colonies, but that takes ~2 weeks. Is there any way to avoid this issue?

More details:

First pcr spans primer/genomic DNA junction

Second pcr amplifes from a tRNA gene that the plasmid disrupts.

Secreted recombinant transient protein expression

My PI tells me that you can't make glycerol stocks out of E. coli cultures grown in super broth. However, I can't find anything to substantiate his claim.

Is this true?

I've made glycerol stocks of E. coli with 2xYT before... which has about half the nutrient content of SuperBroth. So, I would assume super broth would be just fine to make a glycerol stock with.

Thanks

Standard practice for plasmid minipreps is to grow ~5 mL ecoli cultures overnight (~16 hrs). This works well and I always get good plasmids.

However, a few times I wanted just *a little bit* of plasmid. So I only incubated for a few hours:

2 mL cultures for ~7-8 hrs at 37C.

The culture is a bit cloudy by this point and gives an okay yield of plasmid DNA.

However, this plasmid DNA is always seems very poor quality.

If I submit it for sanger sequencing, the results are a little ugly. As if I have a couple SNPs. A tiny bit of contamination. The plasmid is there, but it is dirty.

If I try to electroporate it into bacteria, it always "arcs." Like there is very high salt contamination.

I must re-transform it into ecoli and do a proper, 16 hour overnight culture to regain good plasmid. Then the sequencing is clean and perfect, and it electroporates with no arcing.

Why is this? Why do dilute, low-volume cultures always give such bad plasmid DNA? I would have thought "young" culture would be cleaner.

It is very frustrating because short miniprep cultures seemed so nice and convenient. But the prep is so dirty I think I am wasting my time.

Hello,

If the gene our interest to amplify does not follow any criteria in designing primers how to proceed with that sequence? If the gene sequence is1000 - 1500 bp.

Thank you,

Ramanjaneyulu

My gene of interest cloned in pET28a with C-terminal His tag is expressed in arctic express E.coli strain. Protein is in soluble form and using GE fast flow Ni-NTA beads for purification. My protein of interest is getting eluted in flow through. Please help me in this regard

When I am cloning I frequently see primer dimers instead of the proper PCR product.

T4 DNA ligase is routinely used to ligate two RNA strands using a DNA splint. We are, however, trying to ligate a DNA strand to a RNA strand (using a DNA splint), i.e. the 3' end of the DNA should be ligated to the 5' end of the RNA.

Hello everyone,

I have a vector backbone of 5.5Kb in which I need to ligate an insert of ~600bp. The Roche T4 DNA ligase manual I'm using says ligation should be kept for 16h @ 16 deg. Celsius. My question is that does it really take that long or I can transform the ligation after 7-8h incubation at the said temperature. If not, can something be done to accelerate the ligation?

Note: I do not have the option of using fast ligases that are available commercially. I have only Roche T4 DNA Ligase, which recommends 16h incubation for successful ligation.

Thanks in advance...

Shridhar.

What is the best protocol for the extraction of DNA from avian blood without ordering a kit. If possible please include exact protocol.Thank you

Has anyone had previous success in amplifying Lactadherin from the addgene construct below?

I'm looking at the sequence and it seems that the starting sequence of Lactadherin has too large GC content (83% GC) and annealing temperature (TM= 82'C) for PCR. Please see attached image for the forward sequence.

We perform PCR using Q5 HiFi DNA Polymerase and compute the Annealing temperature based on NEB's online TM calculator. The maximum TM recommended is 72'C.

Could anyone recommend primers that had good success? Thanks.

Which company supplies these?

Hi All,

I am trying to generate a bacterial operon construct. I have designed four gBlocks (each of which is ~1.7 kb) and synthesized them from IDT. I am using pGex-6p-1 vector to assemble these gBlocks using Gibson assembly. I prepared my vector by PCR (primer designed by NEB builder software). After PCR amplification, I checked 5 ul of my PCR product in the gel and found a nice clean single band and the size of this band matches to the expected size (for details see the attached PPT slide). After that, I purified the PCR product using Promega PCR and gel purification kit. I eluted in 30 ul of nulcease free H2O and then measured the concentration of PCR product using NanoDrop and the concentration was 296 ng/ul. Then, I digested 8 ul (296 ng/ul) of purified PCR product with DpnI (1 ul) in a 10 ul of total reaction volume (and incubated at 37oC for 30 mins) as suggested by NEB Gibson manual and heat inactivate the DpnI at 80oC for 20 mins. After that, I set up a Gibson assembly following the instruction of NEB Gibson manual. I incubated the Gibson reaction at 50oC for 1 hr in a PCR machine and then transformed 2 ul of assembly reaction in 50 ul of NEB 10-beta cell (High efficiency) following the transformation protocol for NEB beta cell as specified by NEB. In the following day of transformation, I found many colonies (see attached PPT for picture). I took five well separated colonies and then grew a 5 ml overnight culture. After that, I isolated plasmid DNA using Promega plasmid purification kit and digested them with SalI-HF enzyme which should give three bands if there is a assembled product. Otherwise, they should produce linear vector as pGEX-6p-1 has an internal SalI site. I then checked 60 colonies by colony PCR and it seemed that all of the colonies I checked by colony PCR contains the intact pGEX-6p-1 ( I did not include this gel image in the PPT slide). What things might be wrong with my cloning? Specifically, I would like now

i) Why I am getting intact pGex-6p-1 whereas I generated the linearized the pGex-6p-1 by PCR and digested the PCR product with DpnI? Is digestion not working??

ii) What might be the good way to assemble these four gBlocks into pGex-6p-1 vector?

iii) How can I reduce the vector only background?

iv) I would also highly appreciate any suggestion on my strategy that I am using to generate this operon construct.

For details description of my cloning strategy, please see the attached PPT slides.

Thank you all,

Hassan

I am creating a multigateway vector with the cassettes attB4-attB1R, attB1-attB2 and attB2R-attB3 and I have no problem for the B1-B2 but I fail in getting any colony for the two others. I've been using the B1-B2 as a control in every reaction I did and so far, it worked perfectly each time. I've used different strains and stocks of bacteria, played with the ratio vector/PCR product, re-ordered a fresh BP Clonase II Enzyme Mix and still, it does not work... Can anyone provide me some ideas of things to try?

I want to use the 16s rRNA method. Can I use genomic DNA as a template in this method?

I have been trying to clone a genomic region (Promoter + InEx) that sums up close to 10kb, so far it has been unsuccessful with restriction based cloning and have opted for a gateway strategy which now too has become a headache. I am amplifying my fragment with attB1 and attB2 sites added to my primers and afterwards proceed to purifying from the gel. I have also done the respective calculations to leave the reaction equimolar with the pDONR201 plasmid and left incubating the reaction overnight (up to 18 hours). Afterwards I transform DH10B E. coli and end up with 1 or 2 colonies growing on kanamycin plates if any at all and only after a long incubation!!!! I have also tried a Carbenicillin resistant version of pDONR221 and have only gotten empty vectors which in principle should not happen since there is also de presence of ccdB within the vector?? I have also tested different BP Clonase II batches and have ruled out the enzyme mix.

If anyone has any experience in cloning fragments this size or has had to deal with any of these issues before I would appreciate any input!

Invitrogen discontinued this product. Please let me know if you have some.

Which, do you think, can be the most time-consuming and difficult part of genome editing protocols? Design (oligos, shRNAs, KO, KI, etc.), cloning/construct preparation, screening, etc.?

Hi all,

I recently transduced HEk293 cells (12-well plate setup) with the primary stock virus (0.5ml/well - 1:10, 1:10^2, 1:10^3) to get an estimate of the viral titer. It has been 24h after transduction, but I cannot yet see any GFP positive cells. Though the protocol indicates 48h incubation before fluo assay, I would expect to see at least low levels of GFP positive cells 24h post transduction... Has anyone experienced this before? How long does it normally take before GFP becomes visible in transduced HEK293?

Many thanks

Hello everyone,

I've been trying to perform a ligation for the last couple of weeks and I just get empty plasmids.

The insert is pool of fragments amplified by PCR with an average length of 3kb and cut with the same restriction enzyme on both ends. I am trying to insert this pool of fragments in a vector that is 5,5kb long and is cut by the same enzyme. Not the best option for ligation but this is something I cannot change at the moment.

I am aware that religation of empty vectors is quite probable here so I performed double dephosphorylation of the vector (FastAP and Antartic). I also tried different ratios of vector: insert (1:3, 1:1, 3:1) and I keep getting a similar number of colonies in control and ligation plates.

I've also made some colonies PCR and none of the 30/20 colonies picked by plate showed an insert band. Every clon was an empty plasmid. I am considering moving the protocol to a vector that allows white/blue colonies screening but still, efficiency is really low if there's any....

Does anyone have any advice or suggestion for the ligation? Am I overlooking or missing something?

Thank you very much in advance to you all

Jorge

I have been struggling with a cloning and its due to the fact that It is digested only with a single restriction enzyme(I have no other option). I was wondering why do we need 1 pmoles of DNA for Alkaline phosphatase reaction? Can not it be done with 1 ug of DNA(7kb)?

I generally follow NEB protocol which claims that Restriction enzyme and rSAP can be used at same time during digestion. I did some success with less than 1 ug DNA digestion but this time my bad luck was at its peak. All positive clones had wrong orientation(feel the pain).

I used 4C and 16C incubation with 3:1 and 9:1 ligation ratio. Insert size~ 2200bp and vector size~ 5200bp.

I also heat up ligation mixture at 45C for 5 mins before adding buffer and Enz(it was brought back on ice to cool down ,then I added buffer and enzyme)

Hi everyone,

A while back a lithium acetate transformation of a triple knock out mutant Candida albicans strain was performed yielding several transformants. The transformants, however, integrated the genes at the wrong locus: the genes belong to a multigene family with high homology (~89%). The transforming DNA was prepared using a restriction digestion reaction because the DNA is integrated into the genome via homologous recombination. The protocol that was used to obtain the transformants is currently being used again, but this time no transformants have been identified. Additionally, the cells take several days to grow up (4-5 vs the recommended 2-3), exhibit only a few colonies per plate (2-4 colonies and sometimes none) and consistently exhibit growth on the negative control. (This transformation has been performed several times with the same results every time)

Any ideas as to what the problem might be, or suggestions for how to approach this problem? Any advice is greatly appreciated. Thank you!

The transformation was performed according to the protocol below.

Transformation protocol:

Grow up 4 ml YPD starter culture at 30 deg. C to OD600 ~ 0.8.

Inoculate 50 ml YPD overnight culture with 250 ul of starter culture.

Inoculate 50 ml YPD 4-6 hour culture to OD600 ~ 0.2

Harvest cells when OD600 ~ 0.8.

Centrifuge culture at 3500 RPM for 5 min, remove supernatant.

Wash pellet with sterile water, pellet again.

Re-suspend pellet in 0.5 ml TELiAc.

To two empty microfuge tubes add 100 ul of cells.

Add 5 ul of 10mg/ml ss DNA to both tubes.

Add 5 ug of transforming DNA to one tube, other is DNA control.

Incubate at room temperature for 30 min.

Add 0.7 ml of PLATE mix (LiAc + TE buffer + 50% PEG) to both tubes.

Incubate at room temperature overnight.

Heat shock cells at 42 deg. C for 1 hour.

Centrifuge cells at 5000 RPM for 3 min.

Re-suspend pellet in 0.2 ml sterile water.

Plate on selective media. (300 ug/ml CloNat)

Incubate at 30 deg. C for 2-3 days.

GFP is expressed in B. sbutilis but remains in cytoplasm in spite of secretion signal, even when fused to secretory protein.

In this article fig. 1. The Author has made a decision tree with 24 nodes. Each node is specified for a specific cancer tissue of origin and the couple of MicroRNA which can identify these cancer tissues of origin. My question is, if I isolate miRNAs at the node14(hsa-miR-21, let-7e), node21(hsa-miR-205, 152), node24 (hsa-miR182, 34a, 148), node10(hsa-miR-194, 382, 210), will it be enough to identify cancerous tissue originated from the lung.

Why am I asking this question, because, I want to identify cancerous tissue, which has migrated to different region but originated in the lung. So if I take miRNAs from those specified nodes, will it be enough to identify lung cancer tissue, which has migrated to different region but originated in the lung.

What is the best percentage of glycerol stock preparation for transformed culture is recommended for the long term storage? How long will cultures be efficiency maintained?

I would like to know if there is any protocol or personal experience to cryopreserve chemically competent E.coli cells (for routine subloning) at -20C. The only ultra-freezer that I have access is about an hour driving from my lab. Thanks!

I've been trying to clone a ~100 bp insert into a ~12000 bp vector backbone. Basically, what I've done is to do the standard things (restriction digestion, PCR cleanup/gel extraction, ligation, transformation...).

However, after extracting my plasmid from my transformed Top10 cells, the sequence was very odd. The front part of the sequence seems fine and that's where my insert is. As I progress towards the middle (the vector backbone), I noticed several single nucleotide mutations or SNPs.

I even sent my original vector for sequencing and the sequence looks fine.

What could be the possible cause for this phenomenon?

I have purified my PCR product with PCR purification kit by thermo scientific. When analysing the sample in nanodrop after purification I get a very high absorbance at 230 nm. Will this interfere with subsequent steps like restriction digestion and ligation? The gel purified products alo give me the same problem; absorbance at 230 nm. I use the same kit for gel elution also

I would like to know for how long can I store X Gal and IPTG solutions at -20°C and use it to select transformed E coli without any loss of efficiency?

What are your transfection efficiencies?

I am trying to clone a promoter sequence using a pUC57 plasmid backbone in DH5alpha cells. The finished sequence will contain 6 direct repeats of a ~50 bp sequence I started from complimentary oligo nucleotides with matching overhangs to the cloning site I modified in the vector backbone. Subsequent cloning steps make of two restriction sites with compatible ends to recursively insert another copy of the sequence at the end of the piece already incorporated into the backbone. I have gotten the recombinant plasmid containing 4 repeats very easily. However, on the last cloning step where I ligate an insert containing 2 or the repeats into the backbone containing 4 repeats, I get wildly varying results from colony PCR. Furthermore, when I select the colonies that do show a correct insert size for liquid culture and plasmid extraction, I find by PCR or restriction digest that a deletion always occurs bringing the copy number back down to 1, 2, or 3.

With this cloning strategy I am trying to follow almost exactly the method described for construction of the same sequence in a previous paper, albeit I am using a different vector backbone (pUC57 instead of pBluescript II) and my own restriction sites flanking the sequence. I am even using the same E. coli DH5alpha strain. So I know that cloning and keeping this sequence intact should be possible. I am aware that tandem sequence copy number mutations such as I am experiencing can occur even with E. coli RecA mutants, but I am wondering if there is an E. coli strain out there that would have a chromosomal background more suitable for cloning this tricky sequence. Another concern I have is that after I successfully construct the sequence and after a couple additional cloning steps I will mobilize the finished vector to Agrobacterium tumerfaciens cells. How can I be sure my sequence won't undergo additional mutations in my Agro strain?

One other factor that might be causing a problem is that the repeat sequence I am trying to cloning contains a lac operator sequence. I wonder if it's possible that this is causing some sequence expression problem in the DH5alpha cells where plasmids with 6 copies (but apparently not ones with 4 copies) are at a selection disadvantage. If this is the case is there any techical way to suppress this response in DH5alpha or to use a strain that will be potentially less susceptible to this issue?

Any feedback or recommendations are appreciated. I've run into so many road blocks on this project, I am nearing the end of my rope for coming up with fixes.

Thanks in advance.

I have been currently currently cloning one gene into pTYB12 vector and we had troubles with it all the time.

Anyway, because we got no expression, I was currently trying to delete few first amino acids - putative vacuole targeting sequence. I've sent it to sequencing and the result at first glance didn't work. But when I checked once again, I found out, that the end fits to the plasmid (without the nucleotides as intended), but the first ~770 bp did not align. BLAST found 100% homology to E. coli genome sequences, but also to some weird plasmids, which had alignment at like 5 positions (again 100% identity, no gaps, no anything). When I checked on the internet, those plasmids are probably from E. coli isolated from nature. Some of the positions, where it aligned, were assigned as transposome elements or something like that.

It just seems weird, that it was incorporated just 2 bp downstream from the sequencing primer position, but I guess that's just a chance.

Anyway, my question is, whether has it ever happened to anybody of you, that your plasmid contained some transposome after cloning.

I am working on molecular cloning. I used PCR to check my cloned plasmid. The PCR results show that the primers can bind and amplify the inserted DNA fragment. But when the sequencing results came back, it seems that only part of inserted gene is present. In particular, the part that the reverse primer can bind is lost. I also checked the plasmid using restriction digestion and the bands on the gel are as expected.

I am very confused. Has anyone on ResearchGate encountered similar problem before? Thanks

I am PCR amplifying a 2.8kb insert with the KasI and SacI restrictions sites added, w/ 6bp protector group on each side. I gel purify and then digest this insert as well as my vector by double restricting at 3 hours and 37C (these are NEB time-saver and 3 hours should be sufficient). I then gel purify and ligate the restricted products at 4C O/N. I transform ~5ul into 50uL, or ~10uL into 100uL competent cells using heat shock at exactly 42C. After plating, all my controls are successful (T4 ligase works on single restricted, double restricted vector without insert yields only 1 colony, my competent cells are competent and the transformation of my uncut vector yields 100s of colonies) and the only thing unsuccessful is my cloning plate. I run my ligation mixture and see my ligated product, but I also see a fair amount of insert and vector.

At this point, I've been experimenting for a while and want to give up. But what I conclude from my results is that my insert is not ligating, meaning it is possibly not fully double restricted.

How can I verify I have double restriction of my insert? What else could I be failing to pay attention to? I have been able to verify double restriction of my vector.

Is there something specific to keep close attention to when cloning relatively larger inserts to larger vectors?

Hi all,

We have HiLoad Superdex 200 16/60 Gel Filtration column in our lab, which have some 'Sticky Proteins' and protein aggregates bound to the column. As such, I wish to perform some rigorous cleaning of the column to get rid get rid of these unwanted debris on the column?   

I have done some reading from the column manufacturer manual (GE) and they have recommended 30% isopropanol and 70% of ethanol together with high denaturing agents (Ie. 6M  Guanidine Hydrochloride/8m Urea).

Based on your  experience and opinion, what would be the best approach/cleaning solutions.

Thanks.