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Molecular Cloning - Science method

A platform to ask questions and discuss the procedures involved in Molecular Cloning.
Questions related to Molecular Cloning
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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?
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As I said earlier, if medium or moderate is working for you. It's OK, that is why it is a research but I strongly believe that every good work should have reference point or standard as a guide to you. I will advise that you use what is moderate copy for safety.
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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?
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It looks like your insert fragment is toxic to the E.coli, which selects mutated sequences in vivo. I guess you get fewer colonies than normal?
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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!
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No. It wont. Only in prokaryotic system, where multiple genes can be translated from a single mRNA. Not in eukaryotes. There are other ways it happens in eukaryotes ex IRES-mediated translation. But do not complicate it. The simple approach is to remove the stop codon from the PIWIL4 gene and make a single gene of PIWIL4-EGFP with stop codon only at the end of EGFP. If you want and it is better to introduce a glycine linker sequence between PIWIL4 and EGFP.
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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.
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Hi Balaganesh,
You have probably sorted this by now; but if not, make sure you do not include a stop codon when designing attB PCR primers if you have a C- terminus tag. If you have an N- terminus tag, (ideally) you do want to include a stop codon.
The recombination sites should not affect the transcription of your gene-of-interest.
Good luck!
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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 ?
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I had a similar problem with a protein that was apparently lethal to E coli. No matter what I did there were always indels and my transformation efficiency was super low. Try using a new bacteria strain with pLysS to prevent accidental transcription. Or supplement with 1% glucose in the media. Or do what I did and just screen a looot of colonies.
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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.
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Gelred has a second longer wavelength absorption of around 500nm as well as the stronger absorption maximum around 280nm and an emission spectrum at around 600nm. This makes the blue part of the visible spectrum in normal light conditions capable of exciting the gel red which then produces an emission also in the visible range
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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?
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Hi everyone,
I am just trying to do it with 4 inserts and a double digested vector with BsaI and BbsI. The information about vectors is as below:
1. 208 bps (17 bp overlap with vector)
2. 1254 bps (20 bp overlap with 1)
3. 815 bps (20 bp overlap with 2)
4. 114 bps (25 bp overlap with vector)
I used different molar ratios using NEB calculator:
I tried 1:2 for all
1:1 for fragment longer than 1000bps
1:5 for lower than 200 bps
and different combination of all
1 hour of incubation at 50 celsius. And also used both 10 µl and 20 µl reaction and finally transformed 5 µl of it to the competent cells.
but no colonies...
Would you please help me with it?
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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.
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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."
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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) ?
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Thanks for your kind suggestions, Prof. Murakami~ Shin Murakami
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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.
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Your welcome Olatunde Omotoso.
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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.
1. Synthesise ssDNA commercially:
primersite1 - RE site - DNA sequence - RE site - primersite2
2. PCR with primer1 and primer2 (Phusion)
3. Purify dsDNA product
primersite1 - RE - DNA sequence - RE - primersite2
primersite1 - RE - DNA sequence - RE - primersite2
4. Digest with REs to remove primer binding sites
- RE - DNA sequence - RE -
- RE - DNA sequence - RE -
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Alexander, check your vector. It usually has T7/Sp6 promoter regions flanking the MCS. After cloning your sequences into it, it's easy to amplify them with a pair of these primers.
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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.
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In my experience, calcium phosphate transfection does not work as well as other methods. If you are looking for a cheaper alternative to lipofectamine, you could try PEI (polyethylenimine).
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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?
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Hi Dr. Duguay how big are your overlaps?
I havent tried it yet, but Im planning on trying it out next week. Ill keep you updated...
My plan for getting the fragments out of the yeast was to assemble all fragments in yeast with the YAC or ARS and then PCR out the relevant cassette (-) the yeast stuff, circularize, and then transform back to bacteria
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Dear all, After many clonings my cloning is not working in this pesific vector so I wanted to discuss maybe there is a step that I missed.
I cut all my vectors and PCR products for 1.5 h in 37 degree, I gel purify the Vector and PCR purified the PCR product( I also tried from gel too).
I do use BclI and BsteII restriction enzymes.
Here are my steps and controls.
This is my multiple cloning site agaatcggacgggTGATCAtgctacgcgttgctGGTTACCggtagtctcaa
I modifed vector by mutagenesis and created this MSC. This is the original vector https://www.addgene.org/21894/ I insert my new enzyme sites next to the Fse I site. I use Dam- bacteria for BclI digestion.
I did cut with BclI and BstEII seperately and they both cut the vector in 1h. Sequence change is verified by sequencing.
These are my F and R primers
miR105-5p
5‘AGAtgatcaAATGCACCATCGTTGTCTACATTG 3‘
5‘AAGAGAGggttaaccGGGATTTAAGTCCAAGCAGTGAG 3‘
nearly 300 bp product. and the vector is nearly 6 kb.
I try 1:3, 1:7 ratio with 100 ng vector and I make over night ice-melting ligation.
I transform 7,5 uL of the reaction into 100 uL compatent cells. I also tried 5 uL.
I also transformed 7.5 uL ligation mixture with 50 ng uncut vector to check how efficiency changes. Transformation worked. So there is no problem in transformation step.
My T4 enzyme is new, and it worked in a different restriction cloning.
Honestly, I couldn't find any answer by myself.
Which step could be the possible reason that create problem in my cloning? I always get 0 colony.
Best Regards
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The reason may be that the Restriction enzymes used do not work..the Restriction enzymes can be replaced with others..also it may not be that a substance was used in the medium that stimulates the cloning process, for example, the use of calcium chloride increases the success of the clonation process, as well as hydroxylamine increases the success of the cloning process in Yeasts
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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.
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Because there is no self-ligation problem, there must be other limitations. You could increase the amount of insert and improve ligation conditions. Do you have a ligation control (plasmid digested with single enzyme and then ligated)? This will assure you that everything else is fine except the insert concentration.
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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.
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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!
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The low concentration of DNA contributes, a lot of it is from salt contamination which is common in gel extractions. One way to lessen it is to leave the wash buffers on the column for a few minutes each time instead of immediately spinning, which gives salts more time to solubilize off the column. Another way to lessen it is to just use a newer kit (assuming you are using a kit), I've noticed with gel extraction kits that the buffer which dissolves the agarose starts of clear but goes yellow-orange over time (oxidation probably?) and usually the older the kit the worse 260/230 concentration I get at the end. I don't know if that means the colored contaminant in the buffer just absorbs UV very strongly or if it just isn't very soluble in alcohols and doesn't come off during the wash steps.
It's pricier than ethidium but that isn't too much of an issue for me because I only use it for gel extractions so a single tube of it lasts a very long time. You need a transilluminator which can light up in a blue wave length instead of UV, and either an orange filter or orange glasses to block other emission wavelengths. The illumination happens within the visible spectrum, unlike with UV, so without a filter to block most of it you'll just end up staring at a blindingly bright blue light in a dark room and won't see anything. You can illuminate it with UV but obviously that defeats the purpose. I haven't this myself to know if it actually works but you might be able to cheat with a bright blue LED flashlight and some orange safety glasses in a dark room. I've also noticed if you load a ton of DNA into it you can actually see a visible orange band under ordinary lighting but obviously that's not ideal for most applications.
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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?
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Are you sure it doesn't have rop, and it's not just present but unannotated? Origins of replication and the areas around them are often annotated really poorly because it's inconsequential information for most researchers.
If you are certain that's not the case then I think you would have to give the actual sequence of the plasmid for anyone to really speculate what's going on.
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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!
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Hello, usually small cas9 tags don't affect its performance. If you are worried you can add a self cleaving peptide between your tag and Cas9. We do this with fluorescent protein tags putting different FPs with Cas9 to differentiate clones. Didn't try this in sequencing but i guess it should work.
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Hello !
How can I eliminate primer dimers accumulation in pcr?
Thanks in advance.
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Pre-heat your thermal cycler to 95 degree C and then load your samples. Reduce your primer concentration. Reduce your annealing step duration. Use gradient function of our thermal cycler to obtain the optimum annealing temperature. Limit to 35-40 PCR cycles to prevent unnecessary non-specific amplifications.
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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!
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If DH10B Cells did not work & Stbl3 did, + the fact that you mention " we got a few correct colonies" suggests to me the possibility that your plasmid product may be toxic to E. coli which often causes the bacteria to cleave out the offending product. The Stbl3 is designed to help in those situations. Just FYI, Another strain of E. coli that works well in these situations is called "Copy Cutter".
Ron
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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?
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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.
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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.
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I think you have answered the question then, if it works with the "strong" antibiotic but not the weak one, then you most likely are getting mixed cultures and have background cells that are not resistant. Even well separated colonies can have a small amount of cross contamination, in fact there are likely to be background cells present in an area of the plate without any colonies, unable to grow but not really dead.
Best solution is to either increase the selection pressure or preferably streak out and get pure cultures.
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Secreted recombinant transient protein expression
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I know this post has been here since 2013, but I recently found a platform by DTU (called SignalP) very helpful which I would like to share for the sake of interested learners.
I've come to experience that the efficiency of a signal peptide depends on the nature of the GOI. This is why the SignalP tool comes in handy. Just put in the full amino acid sequence, including the leader sequence, and it will inform you of the secretion efficiency. You can keep changing the SP until you get a score >0.8. Check it out!
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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
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It does not matter that much what you grow them in. What is important is that you add glycerol to at least a final concentration of 10%.
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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.
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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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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.
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John Schloendorn As you said modern methods for cloning work most of the time (which means they actually fail sometime...), just like the old one which has been working most of the time for the last 50 years or so. I do CRISPR/Cas9, I do Gibson and still I use restriction enzymes for some cloning purposes (it's not expensive at all when you are actually using the existing stocks of reagents from the lab!)
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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
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Golla Ramanjaneyulu You can get help in designing primers from NCBI using primer-BLAST. Various tools and softwares are also available for designing primers.
You have to keep following things in mind (you can modify according to your criteria):
1. The primer sequence is annealing to unique sequences that flank the specific target and not to other non-target regions in the sample.
2. Having 18 to 24 nucleotides.
3. Primers having 40-60% GC content of the template.
4. Avoid complementary sequences at the 3′ end of the primer pair. Avoid GC-rich 3′ end. Also avoid mismatches with the target at 3′ end.
5. Design primer having G or C residues in central regions and 5′ end.
6. Avoid sequences which might have the potential in forming internal secondary structures.
Hope it helps.
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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
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May you need to review the composition of your lysis buffers the presence of imidazole could be hindering protein-Ni2+ interactions. On the other hand, the column may just be saturated so you may need to reduce the amount of lysate that you are passing through the column
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When I am cloning I frequently see primer dimers instead of the proper PCR product.
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If you had visible primer dimers that would be an enormous molar excess over full length insert. In that case you would never see the correct clone. Even a small, invisible amount can have enough molecules to compete with your large insert.
The best way to get rid of them is gel purification. Some column-based PCR purification kits can remove primer-dimers too. Look for the size range in the spec sheet. Finally, you can try increasing the PCR annealing temperature to reduce primer dimer formation.
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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.
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Hi, I faced a similar question if it is possible to ligate RNA to DNA using a splint. Five years have gone since you add the question, I wonder how you solved this problem?
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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.
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With NEB T4 DNA ligase (not the fast one), I incubate the ligation mixture for 30min-1h at room temperature and do the transformation. Normally I use 1 vector: 5 insert for my ligations and it works perfectly fine.
Cheers
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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
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@ Abeer A.I. Hassanin
Due to this unique property, very small amounts of avian blood can be used for DNA analysis. Once collected blood can be stored using a range of methods including Queen's lysis buffer (Seutin et al 1991), FTA® cards, in ethanol or frozen whole.
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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.
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Here is what you can try:
1-You never mentioned the tm of the rev primer but looking at the F primer seq
I would suggest to try run a 2 step PCR program at 72 preferably at 70 just to test if you get any amplification (increase the initial denaturation time).
2-PCR with additives (DMSO, or high GC enhancer from NEB) and run at slightly lower tm depeding on DMSO concentration you are adding maybe 67 would do depending on rev primer tm.
3-In case Q5 fails I have found phusion to perform robustly in difficult amplifications.
4-I case all else fails you may want to re-synthesize the fragment and lower the GC content to something more manageable (it is fairly cheap for short length frags).
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Which company supplies these?
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Hi,
If you are still interested in obtaining commercial MSCs, then you can visit this website:
They sell primary cells of various tissue specific and species origin
Hope this helps!
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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
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Simple question, but did you design all of your gblocks with the appropriate overlapping overhangs?
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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?
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I am trying to use this system for the first time. I hadn't realised our destination vector was designed for multi cloning and now I don't have a DONR and the P4 P1r is discontinued. Do you still keep yours? Could I get some? Sorry to bother you but I don't know where else to ask for it...
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I want to use the 16s rRNA method. Can I use genomic DNA as a template in this method?
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Dear all
Genetically, genes are named based on their products/functions. Therefore, there is no gene with the name of 16s rDNA. Also, 16s rRNA sequencing doesn't indicate if the RNA or DNA is used as template. Therefore, 16s rRNA sequencing can be performed based on DNA or RNA templates. Then, you should also check the downstream technique (e.g. genomics/transcriptomics).
I think some authors use the term of 16s rDNA to make clear the ambiguity by mentioning that they have used DNA as template.
Genome extraction ----> 16s rRNA gene amplification/seq. (16s rDNA)
RNA extraction -----> 16s rRNA RNA amplification/seq. (i.e. for gene expression profiling)
Best
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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!
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Thank you for your answer! I will try the separate assembly and hopefully get it to work soon! Cheers!
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Invitrogen discontinued this product. Please let me know if you have some.
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Hello Victor
E.coli DB3.1 is resistant to streptomycin.
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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.?
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I agree.
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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
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Hi Marsha
293T cells is not a good model to determine AAV titer because in vitro cells need very high MOI (multiplicity of infection)of AAV to get a visible GFP-indicated infection, nearly >10^5,with such a high MOI, there was no linear correlation between GFP positive ratio and titer at all. QPCR for WPRE or ITR sequece is a classical method.
Genemedi is experienced in AAV production, you could find more information on this website: www.genemedi.net/i/aav-packaging
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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
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Try using a different protocol. I've easily cloned 7.5 kb fragments with a "Topo-TA cloning kit". No need to do any restriction digests or dephosphorylation treatments. As long as your PCR polymerase leaves an A overhang on the PCR product, you directly add the PCR product to the plasmid vector. The kit is a bit more expensive than most, but it works so well it's worth the cost.
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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).
Would you please share your advice how can I get rid of this problem in addition to my first question?
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)
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The reason that NEB lists pmoles of DNA as the measure is that since the phosphatase works on free ends, the relevant measure is the number of ends, not the total mass of DNA. You can imagine how 1ug of a large plasmid has many fewer ends than 1ug of an oligonucleotide. So the relevant measure is how much enzyme and how many ends (ie terminal phosphates) are in the reaction. You can easily convert ug to pm since you know the mass of your plasmid.
Regarding the second problem of all the inserts are wrong orientation. The good news is that the cloning experiment worked. I would suggest just screening a bunch more, preferably from a different ligation or a different transformation to be sure you aren't just isolating siblings from the transformation. It should just be a matter of screening enough.
However if after finding a lot of clones with only the reverse orientation (that you know are independent clones) then you might have a technical issue where the insert is somewhat toxic in the orientation you want. Probably not likely but possible.
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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.
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Hello, may be you knockout genes which is strictly necessary for grow.
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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.
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microRNAs have to provide potential signature model for various cancers and other diseases. While miRNA in terms of sequencing is difficult , though being done provides huge number of miRNA in a specific cancer as you have exemplified some in your discussion. Few important learning points:
1- Specific sets of miRNAs usually express or otherwise together and they are termed miRNA families.
2-Some miRNA have the potential to act as pan cancer biomarkers but still no specificity is provided.
3-Some biomarkers are like positive or negative acute phase reactant and may rise or fall with non-cancerous disease.
4-There is some but as i experienced little correlation between blood and tissue based miRNAs
5- The science of miRNAs is still emerging and a lot more has to be learnt i guess before they are available, if available for clinical use.
6-Also need to have complete data about pre, pri and miRNA and he cleaving proteins like DROSHA, DICER and factors incorporated in RISC complex.
The whole picture is yet to appear and but hope is there that someday they may be appearing as both for diagnostic use and therapeutic targets like Riversin (spelling ?) for treating hepatitis C.
So potential is there but more research is needed to quantify and deal associated aspects of miRNA
Sorry, that I could not help you straight as i interpret the knowledge about this subject is still evolving.
Kind regards
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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?
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If you had E. Coli glycerol stocks that are repeatedly used. Removed from -80 into a Nalgene labtop cooler that was stored in -80 and returned to -80 in less than 5 minutes. They aren't thawed but still they are subjected to some temperature change. Some of our lab does 25% while others does 40%. Would having a higher glycerol % make a noticable difference in viability over time?
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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!
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Hello, see reference attached.
Only thing after cryoconservation for recovery of culture use fast hitting, just put tube direct to 37 *C water bath.
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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?
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It could be as John suggests, that it is a sequencing problem, which seems most likely for SNPs/ mutations in the vector, or damge to the vector by UV exposure as you suggest.
Before repeating doing anything else, you really need to know if the SNPs/ mutations are genuine.
1)Could these be present in the original plasmid vector - do you have sequencing data from your own lab, or are you relying on sequence data from the manufacturer? I've used plasmids from some manufacturers (no names mentioned) that have several undeclared restrictions sites present, as well as multiple sequence inaccuracies.
2)How long is the good quality sequence read? Check the chromatogram, not just the sequence printout. If it starts looking wobbly of has obvious miscalls or nucleotides that cannot really be called before the SNPs, then the region in which the SNPS are needs to be discounted.
3)If you really want to know what the sequence is then you have options:
-if the sequence is relatively short ( under 6-700 bp and not excessively gc rich or containing hairpins) then you may want to repeat the sequencing with more/ better quality DNA.
-as John suggested, try to sequence using a primer on the other side of the SNPs from the original sequencing primer, or using a primer further forward in your insert, but ideally with area you want to examine at least 100bp away from the primer as the sequence can also be inaccurate in the first 100 bp as well as at the end.
-also if you have a plasmid that has hairpins or supercoils in such a way as to make sequencing of your area difficult, then sometimes cutting with a restriction enzyme that cuts away from your primers and area of interest can make for better sequencing.
Finally, if you confirm that your clone does have multiple mutations in your vector but not your insert then at least you can cut out the insert and reuse, but with short uv exposures.
If you want to avoid uv exposing the vector, then you could use PEG8000 /MgCl2 precipitation to remove short DNA fragments from cutting the multicloning site (https://openwetware.org/wiki/Size_selective_DNA_precipitation). This can also help remove digested/degraded RNA from small scale plasmid preps without columns. It would potentially also work in reverse to keep the insert but not the vector in the supernatant and avoid the need for a gel prep of your digested insert (if small), although I havent tried this myself.
If you are fortunate and have a restriction present in the portion of multicloning site removed from the empty vector, but not in the vector with your insert, then you could add 1ul of the restriction enzyme 5-10 minutes before transformation to reduce the numbers of empty vector coming through.
Sorry for the rambling and hope some of this is helpful.
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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
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I partly agree with Mahesh. The high absorbance at 230 nm is probably due to guanidine isothiocyanate, not ethanol. At high concentrations, it will inhibit further enzyme reactions, but some can still be tolerated. See for example the instruction manual of the Machery Nagel Nucleospin Gel and PCR Clean-up kit, available on line .You can ge rid of most of it by performing a second wash with the ethanol wash buffer and wiping the collecting tube as recommended by Mahesh.
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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?
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1M stock solution of IPTG is stable for 6 months at -20. X-Gal is also stable for 6 months at -20C, however as you may use x-gal vial multiple times it is better to restock it after every 2 to 3 months. Note- X-gal is hygroscopic
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What are your transfection efficiencies?
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Hi Andreas, I usually use H9C2 to simulate the ischemia/reperfusion injury as happened in the heart. You could use adenovirus for transient expression in 14 days or lentivirus for long term expression, both of which can infect H9C2 efficiently.
Genemedi is expert in adenovirus and lentivirus production, you could find more information on this website:
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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.
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Not sure about the Agrobacterium tumerfaciens. For cloning repeat sequence, it is recommended to NEB stable or Stbl3 competent cells. If it still doesn't work, try SURE 2 super-competent cells. Also use lower temperature (30C) rather than 37C for bacteria growth to reduce the chance of recombination.
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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.
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I haven't had it happen personally (that I know of) but I've definitely read about it happening, ex:
If the region that it inserts into is toxic to E. coli then you'll get clonal expansion of the cells with the transposon insertion.
I also know it's happened with some of the larger sequencing projects years ago when they were using BAC libraries for sequencing, I think some of them actually wrote algorithms to find IS elements from the E. coli genome and discard them because it happened frequently enough.
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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
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if you achieved 850 bases in each direction very well done. Everything you are doing is right. Your colleague is right you can fill in the missing sequence using internal sequencing primers. Well done
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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?
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HELLO. Lot has already been said. Would like to add a point here. There are two important points; whether ligation of insert/vector has happened or not? And whether competent cells/strain are efficient enought for such activity?
Confirm your ligated product by using it as template in a PCR reaction using one primer from vector backbone and one from insert. Once you are done with it, focus transformation further. If transformation in e.coli does not work (after changing competent cells/or suitable strain), try to electroporate your ligated product in Agrobacterium cells ( LBA4404, EHA 105 etc). Once it gets stable in it, re-transform to e.coli cells. It works, we confirmed. Good luck
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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.
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Hi,
first with 30% isopropanol 1 CV, wash with 1 CV H2O
- Wash with 0.5 M NaOH 1CV, wash with 2 CV H2O
- additional cleaning using 500 mM acetic acid can be useful. Wash with 2 CV H2O
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Hi guys,
I wanted to know if anyone can help me. Im cloning an 816bp fragment into the MCS of the pk18mobsacB plasmid vector. I have sucessfully cloned the fragment and transformed the vector into E. Coli DH5 alpha (blue/white screening). However my colonies seem to e growing really slow,I carried out colony PCR adn it gives me a positive PCR result when using sequence specific primers for the insert. When I tried to grow the E.coli with the recombinant plasmid in LB plus kanamycin ,it grew slowly again and when i extracted the plasmid with a mini-prep kit, I quantified my DNA using the UV spec . howvcer when i ran it down an agarose gel, there was no plasmid visible. So I ran a PCR to see if I could detect the insert but my PCR was negative>
can anybody help?
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Hello,
I wonder why people call this strain DH10beta. It is a "B", not a "beta":
Also the name "pk18mob::sacB" is wrong. The "::" would mean that the sacB gene was inserted into the mob region. I think the correct name is "pk18mobsacB":
Be aware that the sacB gene renders the cells sensitive to sucrose. Make sure that your medium stays free of sucrose.
Best, Ralf
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Several clones had been produced in the lab before Dolly, including frogs, mice, and cows, which had all been cloned from the DNA from embryos. Dolly was remarkable in being the first mammal to be cloned from an adult cell. This was a major scientific achievement as it demonstrated that the DNA from adult cells, despite having specialized as one particular type of cell, can be used to create an entire organism. Please, the question is that what are the steps for cloning of Dolly sheep?
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Animal cloning from an adult cell is much more difficult than from an embryonic cell. So when scientists working at the Roslin Institute in Scotland produced Dolly, the only lamb born from 277 attempts, it was a major news story around the world.
To produce Dolly, scientists used an udder cell from a six-year-old Finn Dorset white sheep. They had to find a way to 'reprogram' the udder cells - to keep them alive but stop them growing – which they achieved by altering the growth medium (the ‘soup’ in which the cells were kept alive). Then they injected the cell into an unfertilised egg cell which had had its nucleus removed, and made the cells fuse by using electrical pulses. The unfertilised egg cell came from a Scottish Blackface ewe. When the research team had managed to fuse the nucleus from the adult white sheep cell with the egg cell from the black-faced sheep, they needed to make sure that the resulting cell would develop into an embryo. They cultured it for six or seven days to see if it divided and developed normally, before implanting it into a surrogate mother, another Scottish Blackface ewe. Dolly had a white face.
From 277 cell fusions, 29 early embryos developed and were implanted into 13 surrogate mothers. But only one pregnancy went to full term, and the 6.6 kg Finn Dorset lamb 6LLS (alias Dolly) was born after 148 days.
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Can anyone share an experimental procedure for the direct transformation of a bacillus bacterium using PCR product or other non-circular DNA fragment?
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Yijie, which amount of PCR product do you suggest to use?
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I have isolated and identified more than 40 hypothetical proteins from E. coli by using MALDI-TOF and LC-MS/MS. Hypothetical proteins are cloned, over expressed and two proteins are characterized by binding study and crystal study.
Most of the proteins are not forming crystals. It is very hard to make deletion mutant of all hypothetical genes and make characterization of hypothetical genes. I have isolated and purified most of the hypothetical proteins by using chromatography. I would appreciate if anyone could give suggestions and advice regarding the protein identification by using any functional studies.
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I think that is very important a refined bioinformatic study of hypothetical proteins for planning a very detailed set of wet-laboratory experiments
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I am a master student and need to learn about choosing the right cloning strategy. Therefore, I really want to read more books about that issue. Any suggestions?
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Hopefully this will also help you .....
Regards
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I was wondering whether you can express the following:
CMV Promoter - NeomycinResistance - viral2a - mitochondrial signal peptide - GFP - Terminator.
In theory the 2a will self-cleave and you get a NeoR protein and a mitochondrial tagged GFP.
However, the T2a ("viral 2a") leaves a proline attached to the N-Terminus.
Will this mess up the signal peptide, so the GFP will stay in the cytosol? Or will it still work and be imported into the mitochondria?
The same question is asked for chloroplast targeting peptides, ER localizaton signals, and extracellular secretion signals.
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I had the same question and found this Cold Spring Harbor protocol/review of 2A peptides, which comments that 2A generally does not affect the localization of downstream proteins except when the upstream protein is targeted to the ER lumen. 
Design and Construction of 2A Peptide-Linked Multicistronic Vectors
Andrea L. Szymczak-Workman, Kate M. Vignali, and Dario A.A. Vignali
Cold Spring Harbor Protoc; 2012; doi:10.1101/pdb.ip067876
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I am looking for someone willing to send me a little of this 
vector (pBI221) used for plant transient expression (35S::GUS). It's been discontinued by Clontech and is no longer for sale nor available from other databases such as AddGene.
If any of you could provide an aliquot of this plasmid, it will be greatly appreciated (I will pay for the shipment, of course).
Thanks!
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 I did get it, thanks! A professor here in Mexico was kind enough to share some with me. Do you need some Paz Estefanía?
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I have cloned 1kb gene in the pET 22b vector (NdeI/XhoI double digest, 1:10 ratio vector to insert ligation).I have got lots of colonies (~60) in the plate as compared to control (5). I have isolated the plasmid from four different colonies and I got amplification (both T7 and insert specific primers) with 2 of them. But restriction digestion (NdeI/XhoI-10U each, for 0.5ug plasmid) didn't work, only cut vector shifted to the ~6kb. What could be the problem?
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Maybe your colony PCR result is false positive, as u r amplifying from genomic DNA. 
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We're working on cloning for C. elegans, and we're having difficulty with what I assume is plasmid uptake.
We're amplifying out a 6kb segment (vector + rgef promoter + RFP), and a 5.6kb (dip2cDNA + 3UTR + GFP) segment, then ligating them together using Gibson assembly.
After inoculating plates and incubating overnight, very few colonies have grown. and upon PCR screening for inserts, very weak results were obtained.
If the DH5a cells are the issue, are there any other non-methylating bacteria that are good for larger plasmids?
I suspect the size of the plasmid is affecting efficiency. Anybody have any suggestions?
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Hi Cynthia,
do you mind sharing your protocol for DNA purification by low-melt agarose and Beta-agarase digest. Then concentrate using a Millipore DNA Fast flow ultracel Microcon units?
Thank you very much!
Gang
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Does anyone know a method that allows to do X-gal staining very quickly? I stain pieces of mouse embryos and it takes 1.5 - 2 hours. Does anyone know whether a very fast method exists?
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How about Salmon-gal with tetrazolium salts?
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I have been trying to clone a gene of slightly more than 4kb into pLVX vector using single blunt and sticky end ligation (PmaI/XbaI site on gene, SmaI/XbaI site on the vector).
I did ligation using T4 DNA ligase at 16C overnight, at various ratios (1:3, 1:4, 1:5, 1:6, 1:10) with 50ng vector in 20ul reaction.
Then I transform 100ul of competent Stbl3 cells using 10ul of the ligation mix as follow:
->put on ice 30mins after mix
-> heat shock 60 secs
-> put on ice 2min
-> added 300ul LB (w/o Amp) and recover at 37C for 1-2 hours
-> spin down cells at 4000rpm for 5 min
-> resuspend cells and spread on plate
My positive control plate with uncut vector (100ng, about 9kb) usually yield just around a hundred colonies and most of the time I get 0 colonies on the experimental plates.
I have been stuck at this for months. Not sure if it's problem with transformation or cell competency. I've tried making fresh competent cells and did get over few hundred colonies, but no colonies mostly for other plates. Except for once I get a single colony but that doesn't seems to contain my gene. Not sure whether the vector+ insert is too large (nearly 13kb) to be cloned? Any suggestions or advice?
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Same problem here. 4.4kb insert and ~7.5kb pLVXTP, both XhoI/MluI double digested. Dephosphorylated vector, ligated at RT, heat shock transformation and no positive colonies so far.. 
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I did circularization of RNA and then synthsized cDNA using reverse primer. i got a controll pcr using both reverse and fwd primer while failed to get anything when use primers in reverse directions to get full length. I have some confusion to discuss here...
ii use SuperscriptIII in place of MMLV. Can this affect cDNA synthesis?
Is cDNA synthesized from circular RNA will be Circular?
is there any specific metod to foolow in cRTPCR ?
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on which topic you are working..actually i am also working on the same project and got the same problem.