Plasmids - Science topic
Plasmids are extrachromosomal, usually CIRCULAR DNA molecules that are self-replicating and transferable from one organism to another. They are found in a variety of bacterial, archaeal, fungal, algal, and plant species. They are used in GENETIC ENGINEERING as CLONING VECTORS.
Questions related to Plasmids
I was done the 2hr restriction digestion of a plasmid but in gel run no fall out came instead a accumulation of something observed at bottom as in image what could be the possible reason
I inserted my D-product artificial-miRNA into pDONR207 DH5alpha, colony confirmed with PrimerA , PrimerB also with attB1, and attB2 primers. similarly, the plasmid was confirmed with the same primers. but after the LR reaction, I tried to do colony PCR with the same condition as the BP reaction. but I am not getting any results. My previous two constructs were cloned with the above method
I want to linearize a plasmid by SacII, which have one restriction site for SacII. Should I keep it for 3-4hr at 37°C for getting at least 90% efficiency? Suggest me any better idea.
I am doing Pcr cloning trying to clone a 3000 bp pcr product into a 9000 bp pcdh_bec1 plasmid .digestion was done using Xba1 and Not1 ,after gel extraction loaded 2ul on the gel and observed faint band of backbone. for ligation used 2ul(10ng/ul) backbone and 17ul of the insert(24ng/ul). didn't get any colonies I am using stbl2 cells growing them at 30 degrees to reduce chances of recombination.
when i followed the same strategy with the Plvx vector using the same competent cells i observed good amount of colonies although efficiency of competent cells was low.
Can someone help?
someone please explain what is the nature of histone binding to plasmid DNA in CYTOPLASM
1) Does histone bind in form of nucleosome to plasmid? or they float in the cytoplasm as individual entity, and these individual entity is able to bind to plasmid?
2) does histone binding to plasmid occurs in non-specific non-sequential manner?
I have sequenced a clone of bacteria. I got the fastq files back.
What is the procedure for the assembly? What are the differences with human re-sequencing?
Specifically, how can I account for plasmids?
I've been imaging mammalian cells (RPE-1) post-transfection and seen the transfected cells (fluorescent) become static and do not divide, whereas untransfected (non-fluorescent) cells in the same culture do. I've used both Lipofectamine 2000 and 3000 for this and seen the same results for both. I've titred the amount of DNA and lipofectamine, and seen the same no matter how much I decrease it. It is not down to my specific plasmids as I have trialled many different test constructs with different products and observed the same. I observe the cells for five days post-transfection and see no division. So my question is has anyone observed division of confirmed transfected cells? If so, can you recommend reagents or give any advice. Thank you!
Since CRISPR base editor systems typically involve dual plasmids, one providing the Cas protein and APOBEC1 and the other providing the sgRNA, it’s common in animal cells to use GFP protein as a selection marker for the plasmid providing Cas protein, and Puromycin antibiotic for the other plasmid. This often necessitates Fluorescence-activated cell sorting (FACS) in subsequent experiments, which can be costly. Is it possible to use a dual antibiotic selection approach, such as G418 and PurO antibiotics, similar to bacterial experiments, for the sake of simplicity and cost-effectiveness in cell collection? Are there any specific advantages to fluorescence selection in animal cell experiments compared to antibiotic selection?
My supervisor suggested that the bp of PCR products is related to the digestion time. I cannot find anything on the internet to solve my problem. I have a PCR product 500bp. My supervisor implied that I should digest it for longer than the plasmid, why is this?
I would like to use these two plasmids from Addgene. How can I know these plasmids are 2nd or 3rd generation? In addition, these plasmids don't have 5'LTR. In fact, two 3'LTRs are presented. Can the gene flanking by these two 3'LTR be expressed after transduction.
Moreover, can I use these two lentiviral transfer vectors for transient transfections into fibroblast?
Addgene #60903: pHRdSV40-dCas9-10xGCN4_v4-P2A-BFP
Addgene #79372: pHRdSV40_scFv_GCN4_sfGFP_p65-hsf1_GB1_NLS Sequences
I am trying to transfer a group of bacterial strains but they loose the plasmid in the down stream processes. Iam wondering if i can be to transform them in a way that the gene of interest gets incorporated in the chromosome.
I am trying to clone a CRISPR sequencing library. After transformation by electroporation, I picked single colonies and did minipreps, then digested with a single cutter enzyme. Most of my colonies look good, with a single band at the right size. However, some of my colonies have the correct band, plus three additional larger bands. What could be happening here? Could it be concatemers or incomplete digestion? Could my single colonies have two plasmids, one at the right size and one much bigger that's not being cut?
Any advice on what might be happening, and how to figure out what's going on with my plasmids, would be really appreciated. I want to know what is in the plasmid ideally!
I performed electroporation to introduce a plasmid into Agrobacterium and then plated the transformed bacteria onto media containing antibiotics to select for the presence of the plasmid, but subsequent PCR reactions for the target gene are negative
I have tried PEI and lipofectamine 2000 to deliver a GFP plasmid into HEp-2 cells, both showed low transfection efficiency. Does anyone have good experiences in HEp-2 transfection?
I have run PCR and digestion (Not1-hf & Nde1) that gave good bands. The plasmid was PET-28-make123 and the PCR product was franken-flag-dsred.
Then I performed ligation and ran a gel with ligation reaction, plasmid only, and PCR product only control and no bands were present.
How do I find out what caused this?
I will be performing transformation in E.coli Stbl3 strain followed by plasmid extraction. However, the Stbl3 strain is EndA+ strain of E. coli . Can someone please guide me with some alternatives to avoid degradation of my plasmid during or after extraction from transformed E.coli Stbl3 strain?
Note: Following plasmid extraction, the plasmid will be suspended and stored in TE buffer.
I am using 4 different plasmid, each encoding different scFv with similar back bone, The only difference between all the plasmid is nucleotide encoding for scFv seq (Antibody Seq).
When i do transfection in HEK293T, i could see difference in transfection efficiency between all the construct, after 72hr.
Say for Example: Plasmid "A" shows 60% transfection efficiency , Plasmid B shows 45%, Plasmid "C" 30% & Plasmid D shows 10%.
1) The expression of receptor for A,B,C,D are high in HEK293T.
2) None of the scFv induce cell death -In Vitro.
3) The concentration of plasmid, the amount of transfection agent, incubation time of reaction mixture are all same for all the construct.
I hope you can give me some advice on this issue.
We have a tagged plasmid (pcDNA3.1+) where the protein of interest is tagged at both N-terminus (Myc) and C-terminus (Flag). No spacer sequences have been used.
When overexpressed in HEK293 cells, the plasmid is transcribed normally (confirmed with qPCR) and the protein can be visualised with a Flag WB (and is of the expected size). It can also be visualised using an antibody recognising the protein of interest itself. However, when using a Myc antibody, the blot is completely negative. The antibody works well as the positive control (another Myc-tagged protein) can be visualised perfectly.
I have checked and the Myc tag sequence is definitely in-frame so I don't know what the problem is. We have also tried deleting the ATG codon from the ORF thinking that perhaps translation was skipping the Myc tag but this made no difference.
Has anyone encountered similar issues and can give some advice? I am thinking to replace the Myc tag with Ha (I really need the N-terminal tag to work) but I am afraid I will encounter the same problem...
Thank you in advance!
Before transfection the viability is around 98% but exactly one day after I add the plasmid and PEI the viability of my cells decreases so much. I check the viability with the Trypan Blue 100 microliters of it with 100 microliters of cell.
I had extracted dna from E. Colli which shown very low digestion even with Hf digestive enzyme It is suspected that becouse of 1-3 minute kept plasmid with Pd3 during plasmid Isolation caused supercoilling of Plasmid which hindering restriction digestion
I am trying create a mutant plasmid using DpnI digestion. I had put pcr using the mutant primers. But then as I could not do immediate transformation, I kept it to store in -20 degree. Now if I use that pcr after 3 days for DpnI digestion and transformation, will it work?
I am doing transformation with several plasmids into E.coli cells. I have used pUC19 plasmid, GFP plasmid and our expression vector plasmid in different transformations and I have used two different antibiotics, Ampicillin and Kanamycin. I expect colony formation for all transformed cells but cells do not form colonies. They are growing but in a different way, like in the picture below and when I take cells from petri, inoculate into LB Broth with suitable antibiotic and do midiprep extraction next day, I got no plasmids. I am dealing with this problem for one month and I have no other solution anymore. Has anybody faced with this kind of problem?
If my plasmid doesn't have Ecor1 and sal1 but i need my plasmid to have those, I am aware that i can make primers with Ecor 1 and sal1 overhangs but how is that done from the beginning?
Last month I stored a double-digested vector backbone after cleanup/purification at -20 degrees and did ligation with my insert but got a self-ligated product. Is it possible that its overhangs get degraded?
Upon linearization of plasmid DNA for electroporation mediated transformation in yeast, is it necessary to precipitate out the DNA from the restriction reaction via alcohol (With 75% ethanol). Isn't it possible to just deactivate the restriction enzyme via heat treatment and proceed towards transformation using the same restriction mixture?
Hello everyone. I am interested in inserting a large-sized fragment (~20-30kb) into the genome of mammalian cells. What methods can I employ to achieve this? Additionally, how can I obtain the plasmid containing this fragment? Both random insertion and locus-specific insertion are acceptable to me, but I prefer the latter.
I did yeast two hybrid transformation a few days ago and no yeast colonies grow in the DDO+ 225 AbA plates after 3-5 days. OD600, carrier DNA, competent cell, as well as the amount of AD and BD plasmid were correct according to TAKARA manual. Can anyone know the reason why no colonies could grow in the plate?
I'm doing construction eukaryotic expression plasmid vector. I'm not sure if I should reserve or delete the two ITR sequences in pXLG. What are the roles is ITR?
I am trying to determine MICs but I need to express proteins in order to get the antibiotic resistance I am looking for. I have used enriched minimal media, but we are exploring another conditions and MHB seems to be the standard for the type of experiments I am doing. Are there any contraindications of using this media when expressing proteins (not the typical bacterial intrinsic resistance)? Also, my plasmids are Ara and IPTG inducible, and cells are MG1655. Thank you.
How to identify the correct size of a large DNA band (>20kb) after being digested by a unique restriction enzyme?
I have recently digested a large plasmid with a size of 20,208 bp by XhoI, a unique restriction enzyme. I checked the fragment's size by running on an agarose gel 0.5%, 100 V, 1h. In theory, it should give a single band of 20,208 bp, and however, I received a single band with ~3kb. Why and how should I deal with that problem?
Thank you so much for any advice~
I have cloned a gene with EcoRI/BamHI restriction sites into p3XFLAG-CMV-14. The sequencing has confirmed that the gene has been inserted in the right direction. For some reason, I would like to reverse the direction of the gene in the same vector backbone. Is there any method to do it?
Thanks in advance!
Recently I have noticed an uptick of instances where my miniprepped plasmids, when sequenced (using Plasmidsaurus), often give back either 2 peaks or only one peak, where this peak is the E. coli f1 phage genome, even if restriction digests prior to sequencing submission come out clean. I do not observe any other signs of lysis in culture, or plaques in plates.
We are not a phage lab so I am uncertain where the contamination may stem. If you have any advice on how to deal with this problem that would be very welcome.
Thanks in advice,
- I have inserted my gene of interest into a plasmid, extracted it, and confirmed it through sequencing. Now, I want to create a qPCR standard curve. What dilution should I use for it? (Plasmid concentration = 275 ng/µL) I have already prepared dilutions ranging from 10^10 (having 36 ng/µL, resulting in approximately 1x10^10 DNA copies/µL) down to 1x10^1. However, when I attempt to confirm values below 1x10^7 using a Nanodrop, the instrument cannot calculate the value accurately, and there is occasional variation in the readings (sometimes increasing and sometimes decreasing).
- When I perform qPCR using the dilutions (ranging from 1x10^7 to 1x10^1), the threshold levels do not consistently reach uniform values. Why is this occurring?
We are currently trying to transform Bacillus licheniformis (environmental isolate) with plasmid pWH1520. Unfortunately, we barely get colonies growing on our plate regardless of the transformation protocol we used. We started with protoplast-based transformation before finally using electroporation but it didn't make any difference. Still no colonies growing and we have no clue why. We also read that some strains might be difficult to transform because of endogenous restriction system that rejects foreign DNA and that transforming B. licheniformis with unmethylated DNA can circumvent the problem. Do you think it might be the reason behind all those failed transformations or are there any critical steps we missed and therefore we might need to incorporate in our current protocol to make it work? Your feedbacks and suggestions are very much appreciated. Thank you.
I 'll design the large sequences (approximately about 1000 bp) as the homology arms.
so for transformation, which ones are better? the linear integrative plasmids or linear PCR cassettes?
I have applied Lipofectamine 3000 and Versatile DNA/siRNA transfection reagent jetPRIME® to transfect PX459 plasmid into A549 and NCI-H1975, all the operations are performed according to the manufacturer's instruction. But 24h after transfection, more than half of the cells died, Moreover, after adding 1ug/ml purinomycin, most of the cells died within two days, and finally the monoclone could not be screen. I'm not sure if it is a problem with my cell activity or something else, here is a picture of my H1975 cells before transfection.
I transformed a library into stbl3 competent cells following a standard protocol which is like incubation 30min in ice, 42°C for 45s, soc culture at 37°C for 1h, before culturing at 30°C on LB Amp+ dishes.
some supplementary info:
1. The dishes look normal after 30°C culture.
2. The plasmid was tested and had no problem.
3. I used another plasmid to tansform and harvested bacteria directly from the dishes(didn't frozen in refrigerator) with correct plasmid.
I can only find one reason for this phenomenon is that -80 made it happened. Can you help me find other factors?
I need to address my protein in the ER (endoplasmic reteculum), for this I need to construct a new plasmid: does the peptide signal sequence need to be added inside my protein sequence (after the N-terminal start codon) or can I merge it just in front between the promoter and my protein sequence ? (see photo).
I have an overnight culture in 2XYT broth-medium, which has been transformed using M13 phages as vector.
I would like to extract the plasmid for gene sequencing. Could someone tell me how long they could be stored at 4°C and if the plasmid would still be fit for sequencing.
Thank you in advance.
Hi, I am wondering if different plasmid extraction kits yields different quality of extracted plasmids, which subsequently affected some downstream analysis such as nucleofection in human embryonic stem cells. I am asking specifically for the used in human embryonic stem cells context.
Could you also provide your preference and the reasons?
Sgv was constructed and maxiprepped. The conc was 1965 ng/ul and accidently a few drops of nuclease free water got spilled in it.
Can I still use the same SGV plasmid for transfection or should I start again?
I was going to run restriction digests on them.
I am currently trying to do a CRISPR/Cas9 knock-in and am wondering about the design of the donor vector (with the HDR template).
I've been looking through many publications, but never actually find an explanation how the backbone is chosen. If the vector consists of an ORI and an antibiotic selection marker, can you just take any backbone, like pUC19 or so on, or does a knock-in require certain plasmid designs (apart from the insertion cassette, of course)?
I have an insert size of 900 bp, trying to clone it in pET28a vector. But after every the transformation when i am doing colony pcr and digestion check i can only get the fragment of 500 bp. Why is it so? The restriction sites are BamHI and SacI and the gene doesn't have internal sites of these, which i could know by sequencing. So why is it so. Do i need to subclone instead of directly trying to clone in pET28a plasmid?
I have seen in many papers up to 8 kb Plasmid DNA can be transfected in mammalian cell line with Lipofectamine 3000 LTX and Plus reagent. But my question is can I transfect plasmid with a size of 13 kb?
Recently I performed an yeast transformation and I got a few colonies in the selective plate. The strain is very modified, with four auxotrophic markers (CRISPR) and we are struggling to insert plasmids in It. This specific transformation was made to insert a third plasmid in the yeast
I screened the potentially positive colonies by colony PCR and I saw no bands, whereas I got amplification of fragments of interest for the other previous plasmids. What would be the problem?
The reaction? Maybe a contaminant containing the same marker selection as my desired plasmid?
The primers for this reaction have the same Tm as the other ones that were successful. Do you guys have any suggestion? Thank you.
Does eukaryote DNA extraction need to perform in sterile condition with sterile (TE) buffer?
And what about plasmid extraction?
I read somewhere that plasmid DNA extraction does not need to be sterile as plasmid DNA is supercoiled and remains in solution when sodium acetate is added. This causes proteins and chromosomal DNA to precipitate and allows for the extraction of the purified plasmid DNA.
I'm having trouble cloning a 700bp insert into a 18kb plasmid. I'm using Neb Hifi assembly (Gibson assembly) but everytime I have no colonies on the plate. I tried many different insert/plasmid ratio but nothing change. I'm using XL10-Gold bacteria. Both insert and plasmid were purified on agarose gel.
Thank you so much.
I have recently been considering the creation of a new competent E.coli strain for further research. I'm interested in incorporating a plasmid into this strain(~10Kb in size | replication origin is p15A ori | express 4 proteins controled via pBad system | Chloramphenicol resistent).
However, I am concerned about potential issues that might arise when I attempt to transform it with another plasmid that I need to extract(~5.2Kb in size | pUC/Origin | AmpR). I aim to extract this plasmid as purely as possible for mammalian cell transfection. I'm worried the incorporated plasmid may contaminate my plasmid extraction process.
While I could attempt to make the bacteria lose the ~10Kb plasmid by withdrawing the antibiotics, I believe this will be a time-consuming process. I've read in some articles that it might take approximately 30 generations for a bacterium to lose a plasmid.
There are two questions I would appreciate some input on:
1. When using a competent E.coli strains such as Rosetta (which includes pRARE) or LEMO21 (which includes pLEMO) to amplify plasmids, is the potential contamination from those innate plasmids a consideration?
2. I am wondering if it might be possible to use DNA size selection methods such as magnetic beads to extract only the smaller plasmid, which is approximately 5Kb in size.
I have received the sequence of plasmid, I aligned it with the gene sequence on APE plasmid editor and it is matched. But when I performed vector screening on ncbi, the sequence is strongly matched to vector. Can I make probe with this plasmid by cutting with relevant enzyme??
This plasmid is quite old and has been stored at -20c since 2008. When I miniprep it from liquid cultures of about 5ml, it always runs slower on a gel, including the digested fragments. I also find I can't generate retrovirus using it whilst I can with the original stock. Does anyone have any ideas as to why this might be? Many thanks!
Please see attached image - numbers indicate different minipreps, OG refers to the original plasmid, and Un to undigested.
I have a insert of 400 bp cloned in vector pbluescript II KS (+) of size 3.0kb at RE sites XbaI and XhoI. But when I try to double digest the plasmid it is not happening. I am sharing picture of result showing the same. Please can anyone provide me the reason and solution for this.
Fig: Lane1: 100 bp plus ladder; lane 2: plasmid double digested; lane 3: plasmid single digested; lane 4: uncut plasmid
I have a large plasmid (20kb) which I am trying to transform into chemically competent commercial EHA101 cells. I had success with transforming a 16kb plasmid into the same strain, but have been unsuccessful with my larger plasmid.
This was my method:
1) Thawed agro cells from -80 in hand
2) added 2.5ug (5uL) of each pGE013_Upf1sgRNA_1
3) 30min on ice
4) 5min in liquid nitrogen
5) 5min in 37 water bath
6) 5min in ice
7) Add 900uL of YEP
8) Incubate at 28C with shaking for 7 hours
9) Centrifuge at 7000rpm and remove 900uL of supernatant
10) Resuspend cells in remaining supernatant
11) Plate on YEP+spec and YEP +spec +kan
Wrap with parafilm and incubate at 28C. Saw growth for 17kb plasmid (binary CRISPR/Cas9 vector) after 4 days on YEP+kan+spec. These colonies grew on YEP+spec+kan+rif. I also purified plasmid from these via alkaline lysis and saw it present on agarose gel but no growth on the 20kb plasmid plate (pMpGWB337 vector with insert).
I had previously tried a shorter incubation of the cells+plasmid and a shorter outgrowth period.
I don't have the materials for electroporation, so I am very hopeful that I can somehow make the freeze-thaw method work.
I transformed a plasmid which self-expressed mCheery fluorence in cells, but it seems to express weakly in E.coli. Therefore, the FACS could not seperate the cells with and without plasmids well. These pictures are cells with plasmids and the blank control (cells without plasmids). I wonder the reason of this problem or if there are methods that could enhance the mCheery fluorescence?
I am a novice in molecular cloning, and recently I was given a new project to overexpress lncRNA in HEK cells. I am looking for suggestion and guidance for the following queries:
1. On what basis should I select the plasmid for overexpression?
2. The length of the gene is 4.2kb. Do I need to split the gene in 3 or 4 parts and perform cloning?
3. Does the size matters for over expression?
Thank you in advance!
Unwanted base pair insertions/detetions!
This is an age old story with this cloning journey for one particular plasmid ~15kB with a CMV promotor. I get unique base single base pair deletions/insertions that I am pretty sure are happening during bacterial replication, not as a result of a bad ligation, etc.
-E.Coli DH5a competant cells and heat shock transformation.
-In several stages of cloning, of 6 screened bacterial colonies, all differ by base insertion/deletion, different location in every clone, breaking my reading frame and rendering the construct useless.
Most recently, I screen a clone (12) and confirmed the sequence of my cDNA is 100% correct and there is just 1 moleular species present.
I want more material, so I go to my agar plate with the streaked bacterial colony of clone 12 use this to innoculate my LB + antibiotic. I grow for 2 days at 30' as it the plasmid is large (15 kB) and I worry a bit toxic to the cells.
After miniprepping this culture, sequencing reveals a new base deletion in my protein that didn't exist in the original parent cDNA.
-Any tips? I have tried growing at 37' for 16 hours, 2 days for 30', leaving it a room temp for a day to start and THEN putting in 30', but I'm not sure what else to try and need to produce more of this construct one way or another.
I am looking for advice from experienced individuals regarding the optimal conditions for plasmid electroporation in A375 cells. Would anyone be willing to share their expertise on this matter?
Additionally, I attempted electroporation using 140, 180, and 240 volts for both 70 ms and 35 ms, but unfortunately, the cells did not survive. Any suggestions on how to improve the outcomes would be greatly appreciated."
I have transformed around 1000 ng of my circular plasmid into E. coli BL21 (DE3). However, there was no colony but a transparent bubble-like 'colony' formed at the bottom of the plate (As attached) . May I know what is it and how to overcome this?
I think there is nothing wrong with my competent cells as the transformation of positive control (blank vector) was successful.
I used 0.5 ug plasmid. Cell confluency around 60%. Transfection time is 48 hour. I used empty plasmid (pCMV-3Tag-4A) and only Lipofectamine as a negative control. However, I used EGFP plasmid tag for positive control. I found Myc tag plasmid transfection rate is only 5 to 10% where EGFP tag plasmid transfection rate is 60 to 70%. I am wondering why it low ? Can any one explain vector pCMV-3Tag-4A and pEGFP-C1 both have tag but myc tag did not express but EGFP is expressed. Is it functional and non functional peptide ?
- I have been preparing competent dh5alpha cells in the lab with good competency not excellent. however, have not been able to transform my CRISPR plasmid yet. I am following all the desired steps still unable to attain the correct colonies. plz, throw some light where I can be making mistakes. Plasmid is from addgene (pSpCas9(BB)-2A-Puro (PX459) V2.0)
I did a transformation of the BL 21 cells with my plasmid DNA. Then I ran the dna plasmid pcr to check the inserted dna and both samples showed bright, clear bands. However, when PCR with a larger volume was available for sequencing, one sample was very faint (with increased plasmid volume) and one sample had many bands. Is the first result a false positive?
So I am performing a routine site directed mutagenesis of a ~ 7kb plasmid. After PCR I DpnI digest my template DNA and then transform my PCR product into DH5a cells. I use a partially overlapping primer design that works really well--typically I don't even bother to optimize my primers as this basic design strategy has worked for me to make hundreds of mutants. Every so often I get some sort of weird primer stitching PCR product, but I haven't really wrapped my brain around how I can get a final circularized plasmid like the one I am sharing.
Thanks for any insight!
Hi, currently I'm working with E. coli Bl21 (DE3) pET-26b(+)-N20-aiiA-6xHis. I'm trying to improve the secretion rate of the aiiA to the extracellular fraction.
"pET-26b(+)-N20" already provide the signal for the extracellular secretion. so i thinks its normal to have thick band in the SDS. but how about the native protein? why it also looks thick? especially the one in the sample are more thicker, than the one in the control.
Both sample and control are undergo the same process. The difference only in the plasmid (control doesn't have plasmid).
thank you in advance!
I am facing a problem to circularize a particular bacterial plasmid sequence while doing hybrid assembly in unicycler. After the assembly, it's not showing circular. While the other plasmids have circularized in unicycler after doing the hybrid assembly. Please give me a solution how that particular plasmid can be circularized during hybrid assembly?
(Text in bold was the error conducted)
Mutation: WPD to FPD
Restriction site: BspEI
Template size: 7.16 kbp
The following reagents were added to make a total volume of 50uL:
5x PCR buffer: 10uL
Primer mix: 2uL
Phusion polymerase: 1uL
template 60ng: 10uL
The experiment method wanted 5ng/uL plasmid DNA solution to add 50ng of template DNA to the reaction. However I made an error in dilution and resulted in 6ng/uL, but still added 10uL as indicated above.
Would or How would this affect PCR results? Thank you in advance for your help.
I would like to transfect HEK293 with a plasmid containing a luciferase (downstream of a response element to this pathway) to check a cell signal pathway, but in order to have a population of cells homogeneously transfected I decided to include a fluorescent protein and stop codon with its own promoter so I can sort cells post-transfection.
My question is...based on the orientation of the plasmid map: response element-Luciferase (Renilla)-SV40 promoter-Fluorescente protein (mRFP1).
Could it be safely used? Do you think that the position of the Luciferase and fluorescent protein are far enough so when I perform the luciferase assay there won't be any light that may interact/excite the fluorescent protein (I'm trying to avoid any BRET)?
Thanks in advance for all the help that I could receive!
I am currently working on making mutations in my plasmid conducting Quikchange. The plasmid size is 3800bp and I am using primers with several mutations. However, both my parental plasmid and the mutated product have two bands (see attached gel)
The gel shows L1) GeneRuler 1kb ladder, L2) my parental plasmid from a mini prep, L3) the plasmid product (mini product) after conducting PCR -> DpnI -> PCR cleanup -> Transformation -> Mini, L4) the plasmid product (mini product) using the mini product from L3 as template for a new round of PCR -> DpnI -> PCR cleanup -> Transformation -> Mini.
It looks like I get less of my desired plasmid for each round (~3800 bp) and more secondary product. The loss of primary DNA is not feasible, as I need to conduct three consecutive rounds of mutation PCR. Can anyone help me explain what the secondary product may be? I have never seen it before.
The lab works with M13 phages containing ~6400 bp ssDNA. Could the upper band of L2 be the plasmid integrated into phage genome (~10,000 bp total) from previous experiments? And could L3 and L4 be the product of more plasmid integrated into the phage genome? Or is this genomic DNA contamination?
I hope that you can help.
Kind regards Emil