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Questions related to Molecular Biological Techniques
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13 answers
I would like to make primers for my qPCR. For that, I need a reliable software. I know the (ABI) Primer express software(paid) is preferred, but do you know any freeware that does the job well?
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You can use the same rules for qPCR primer design as you do for end-point; just shoot for amplicons between 75-150bp if possible.
Also, make sure you validate your new primers after they arrive by performing a standard curve using template that is similar to your experimental samples to determine the dynamic range.
Any of these are nice:
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21 answers
I am designing my real time PCR pair of primers. I put them into oligoanalyzer to determine hairpin loops, homo and heterodimer formation. What are the limts for a good design regarding these 3 items? The program gives the Gibbs free energy for these three posibilities,There are a number beyond which it is not convenient to synthesize? I understand that any program to design primers get a list of the best primers, but taking into account only the specificity, is that right? So I have to decide also to base on how they act about this kind of other possible reactions, right?
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Also design primers that have "span an exon-exon junction".
This controls whether the primer should span an exon junction on your mRNA template. The option "Primer must span an exon-exon junction" will direct the program to return at least one primer (within a given primer pair) that spans an exon-exon junction. This is useful for limiting the amplification only to mRNA. You can also exclude such primers if you want to amplify mRNA as well as the corresponding genomic DNA.
Copy Pasted from:
And Never trust 100% to any Primers from Primer-Bank or website or Papers.
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18 answers
I used to isolate my RNAs with Trizol, purify them with qiagen RNeasy mini kit and on-column DNase digestion before performing reverse transcription and real-time PCR. Unfortunately it was not possible design primers including intron-exon gaps within their target region to avoid genomic amplification. I performed RT with high capacity transcriptase (applied biosystems) kit. My problem is, in qPCR, I still have amplification in RTnegative (no RT enzyme). I tried to increase digestion time and add higher concentrations of DNase. I also added a final step with RNase in RT reaction, but still I'm getting the amplification in negative control RT reaction. The qPCR negatives (water) are clean. Can anyone help me?
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Assuming you are not facing contamination, there are more efficient DNases. The best one for this purpose is shrimp nuclease from Arcticzymes, which is dsDNA specific. You can therefore add more of it, without damaging your primers. The DNase is then inactivated by heat (http://www.tataa.com/products-page/quality-control/).
There is another more cost efficient strategy called ValidPrime that was developed by Henrik Laurell. ValidPrime measures gDNA contamination and assay sensitivity for gDNA and qPCR data are easily corrected for the contamination, if necessary. The advantage is you don´t have to measure RT- controls, which saves a lot on reagents (see same link as above and the ValidPrime paper on www.tataa.com).
Be aware, designing assays across introns does not guarantee you will not amplify gDNA! This is common misconception. Many eukaryotic genes have pseudogenes, occasionally hundreds of them, and they lack introns, because they are copies of the transcripts. Therefore, you must always check for gDNA contamination even when you can design assays across introns.
Good luck!
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18 answers
I am doing induction with Tet-on system. However, both of the tetracycline and doxycycline solutions I used failed to act as an inducer. I don't know how old the solutions are, but I'm suspecting that they are too old to function. Does anyone know how long they will last if stored in dark at -20°C?
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Powder
• 1 year from date of receipt under proper storage conditions.
Solution
• 4 weeks at 4°C
• 3 months at –20°C
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12 answers
I have a problem with plasmid extraction by column. I am trying to extract plasmid by phenol-chloroform. I got the following protocol from nearby lab:
1. Resuspended bacterial plasmid with TE+RNAse (10mM tris-HCl, 1mM EDTA, 100mg/ml RNAse).
2. Lysed cell with TENS Buffer (10mM tris-HCl, 1mM EDTA, 0.1M NaOH, 0.5% SDS).
3. removing debris by mixing with 3M NaAcetate and centrifugation @13,200 rpm for 4 mins
4. purified with saturated-phenol (centrifugation @13,200 rpm for 4 mins)
5. purified with saturated-phenol:chloroform (ratio 50:50) (centrifugation @13,200 rpm for 4 mins)
6. purified with chloroform (centrifugation @13,200 rpm for 4 mins)
7. precipitated DNA by absoluted Ethanol (centrifugation @13,200 rpm for 2 mins)
8. washed DNA by 70% Ethanol (centrifugation @13,200 rpm for 1 mins)
9. resuspend DNA pellet with SDW
-------------------------------------------------------------------
From this protocol, I have some questions:
1. Which step remove gDNA out from plasmid? Is that step3, or not?
2. Can I used the plasmid from this extraction in cell transfection?
3. How long is time for stocking phenol:chloroform mixture? (I've read some question in RG and the answer seem to be within 3 month. So, I don't know about the reason: denatured? reaction?)
4. I got 260:280 ratio from this extraction more than 2, can I used that plasmid? (or that mean it contaminated?)
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Hi Wareerat,
I wonder about that protocol a little bit, because you use 3M sodium acetate instead of potassium acetate. Using potassium acetate instead of sodium acetate is very important, because potassium ions bind SDS and lead to precipitation of SDS and the bound proteins (which sodium does not). Did you observe a lot white precipitate immediately after addition of the buffer at step 3? If not, then the wrong use of sodium instead of potassium might be the reason.
As Tomasz already pointed out it is also very IMPORTANT NOT TO VORTEX the samples, because gDNA is coprecipitated by DNA-bound proteins. If you vortex the gDNA is sheared on the one hand and proteins are detached from the gDNA to a great extend on the other hand, which will lead to gDNA contamination.
The 260/280 ratio higher than 2 indicates contamination with RNA. Addition of new RNase A would be helpful, because degraded RNA will not be precipitated under conditions used.
Best,
Kai
Protein smear in non-denaturating polyacrylamide gel after silver staining.
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polyacrylamide gel "5%". The single strand appears very well, however, the protein appears like a smear. I have
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I think you have to load less protein.. The sensitivity of silver staining is 0.6–1.2 ng.
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9 answers
When I am comparing my 32P nucleotide-labelled DNA sample with a 32P end-labelled maker on agarose gel electrophoresis, the radioactive signal from the maker is much lower. Is there a way to label the marker with radioactive nucleotides as well, in order to increase the specific activity of the DNA fragments?
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Hi Guus,
I don’t think this is an image manipulation only two exposure times and you should indicate clearly that to reviewers and in figure legend. Maybe you should indicate why you have chosen to perform 2 different ways of labeling. The only possibility is to end-label all and have longer exposure time for all. I think you could try that while you are finishing your manuscript. Good luck with it!
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22 answers
I have recently cloned some mice which is an expensive endeavor. The construct I was given to clone has our gene of interest tagged to EGFP on the c-terminal end. My job was to put this sequence into a new vector with a more specific promoter. The construct I used happens to leave the start codon for the EGFP intact - note, the stop codon for our protein sequence has been removed and the EGFP stop remains.
Since that is how the construct was sent I assumed it is okay. Now we have (expensive) green mice, and I am beginning to fret about whether or not leaving the start on could cause problems. Is it okay that the start is still there? Is there any chance the GFP will be expressed without the protein being fused?
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It completely depends on the fusion mRNA. But my gut feeling is that it is unlikely your GFP start codon would be used. Try the western blot as suggested above (Abcam #ab290 works well), but be warned that it is very likely you will find an unfused GFP band by western blot at high protein amounts and long exposures -- especially if you transfect your cloning plasmid. The reason is...
...although one ribosomal subunit accesses the mRNA from the 5' end, translation initiation can occur at any AUG (in any frame) along the length of the mRNA. That said, 5' AUGs are more frequently utilized than AUGs farther 3' (due to the higher frequency of Kozak sequences toward the 5' end and also scanning across other AUGs after initiating translation upstream; an exception to this is translation reinitiation where a ribsome starts at a second AUG after termination). In many instances, usage of downstream start codons is correlated to copy number of mRNAs, suggesting it's a probability game.
Alex Kochetov has done some nice work on this in recent years.
All this to say: In future projects I would think it is probably better to get rid of the GFP start codon so it will only be expressed as a fusion and you won't have to worry about alternative translation initiation mechanisms. Just be as careful as you can with the mouse you've already made. Maybe try localizing the fusion protein by IHC, though I'm guessing that's challenging and is why you made the GFP fusion in the first place.
Best of luck. Shoot me an email at jonathan_lowery@hsdm.harvard.edu if I can be of any help.
Jonathan
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9 answers
I'm extracting genomic bacterial DNA in reasonably large quantities and I need it to be high quality too.
The final step in the protocol I have been given incubates overnight in TE with 0.5% SDS, and 0.5mg/ml each of RNAse and Proteinase. Will the proteinase damage the RNAse?
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Another reason for the two step: TE contains EDTA which sequesters Ca2+ and Ca2+ is needed for RNAse whereas proteinase K can function independently from Ca2+ to a degree...
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2 answers
Does anyone have experience with pulse field gel electrophoresis (PFGE) protocols to determine chromosome number of filamentous fungi / yeast cell protoplasts? I found different protocols but I'm unsure about electropheretical set-ups (voltage, running time, etc.). How it is possible to discern co-migrating bands?
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Sonnenberg, A.S.M., De Groot, P.W.J.,Schaap, P.J., Baars, J.J.P., Visser, J. & Van Griensven, L.J.L.D. (1996) The isolation of expressed sequence tags of Agaricus bisporus and their assignment to chromosomes. Appl. Environm. Microbiol. 62, 4542-4547.
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13 answers
We have been using the SMARTer RACE kit from clontech, and gotten both an expected full length product and a truncated product. However, I am unsure if this truncated product is real because although it has a stop codon, it lacks a UTR (...ATG/AGT/TAA/AAAAA...) and in the longer RACE product the corresponding region has a short polyA rich stretch (...ATGAGTCAAAAGAAAGGAAAA...). I am thus not sure if the adaptor primer is picking up on this short A rich stretch. I have had this happen before myself but usually without the stop codon so I ignore it. Thanks for any advice.
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Hi Jonathan,
Another way to perform 3'RACE without using an oligo(dT) sequence is to ligate an adaptor to the 3'end of your RNAs and use that adaptor to primer your RT. Then you just need to do a PCR using a forward primer which is specific to your gene and the reverse primer specific to the adaptor. This way you don't have any bias caused by gene encoded poly(A) stretches.
In our lab we use T4RNAligase2 to ligate a DNA adaptor to the 3'end of RNA and it works well.
Otherwise, you can always go to the classical northern blot using two probes, one upstream of your putative alternative 3'end and a second one downstream. If your 3'RACE result is correct, you should see two bands using the probe upstream the putative alternative 3'end (one corresponding to the full-length and the other to the truncated RNA) and only one band when you use a probe downstream the 3'end (corresponding to the full-length RNA).
All the best,
Emiliano.
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3 answers
Does anybody have experience in using the dolomite microencapsulation system (http://www.dolomite-microfluidics.com/en/new-prods/droplet-systems)? I would like to embed single microbial cells (E. coli) into microspheres of approximately 20-30 micrometers for subsequent FACS analysis. Has anybody here such a system and used it for generating such bead-diameters? How much optimization is needed before getting such a system "up and running"? Any suggestions of other potential systems are welcome.
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Hi Lars,
Have you had any luck with encapsulating cells yet? If you share your email, I could send you further information. You could alternatively send me an email at ashok.sinha@dolomite-microfluidics.com
Within a day you could start producing droplets reliable and at high frequency. Encapsulating cells is sometimes tricky and mostly related to their fragility.
Thanks,
Ashok
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3 answers
I have a question concerning a commercial Taqman assay, which has been designed for the simultaneous detection of the target of interest (FAM-TAMRA probe) and an internal control (JOE-TAMRA probe). For the positive control, the manufacturer states that the Ct value must be lower than 36, which is the case (32) on our StepOne device (48 well plate, 25 uL per reaction). The internal control yields an average value of 24.5 in our experiments while manufacturer data, obtained with different types of samples, vary between 24 and 26. In our hands, the interassay CV is very low between experiments, both for the positive and internal controls.
The same samples have been tested on another cycler (LightCycler 480), yielding identical values for the internal control, but lower Ct values for the positive control (28). During this experiment, 10 uL/well was used (384 well plate). One could state that PCR efficiency is different between both devices, but this would result in different IC values too (or at least I suppose this). Am I correct? Importantly, our StepOne has no filter for the detection of TAMRA. Can this be an explanation for the different results?
What is the impact on the assay outcome if TAMRA fluorescence is not measured during PCR reaction?
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Hi Frank, for both Taqman probes the Tamra dye is acting as a quencher to the dyes you are actually measuring. Therefore you do not need to measure on that wavelength. The differences between the machines might arise due to different excitation and emission filters for the two machines, thus you might be measuring one of the dyes at a different part of its emission profile. Have you checked the efficiency of the positive control reactions on both machines? The lower volume might have subtly altered the PCR reaction.
Hope this is of some help.
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6 answers
The minimum temperature we can prepare in our lab is -20°C.
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The best quality long-term maintenance of microbial cultures is certainly cryostorage in liquid nitrogen (culture medium containing 10% DMSO or 10% glycerol), if you have this possibility. Storage of yeast cultures at -20 C may be detrimental to them, better is routine passaging.
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I want two of my genes of interest to be synthesized over there, please share your experience.
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We use GenScript for our Taq and dNTPs. I've been very happy with them.
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We aim to assemble 6 PCR fragments (300-400 bp) by Gibson assembly. We assembled and PCR amplified the first 3 and last 3 fragments with no problems. However, the assembly of the two amplicons to the full-length product fails and PCR analysis shows that fragment 5 and 6 are faulty. We are using the NEB Gibson assembly master mix essentially according to protocol. We assume that secondary structure during assembly of the two amplicons at 50 degrees C are causing the problem. Do you have any suggestions on how to solve this problem?
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I completely agree Karl and Abdelhalim. DMS0 (as suggested by gonzalo could be useful too).
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I am trying to measure glycerol consumption in a minimal medium where I am growing Mycobacterium smegmatis.
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Try quantitative LCMS if possible. It will be very accurate.
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8 answers
My genes are more than 1 kb (one is 1.09 kb, and the other is 1.7 kb).
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Genewiz Inc (NJ, USA) is a leading company on gene synthesis. Long sequences are provided for a good price. Take a look!
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4 answers
In case somebody is not able to amplify a gene from a plasmid using a designed primer from NCBI which is not working on clinical isolate strains. Will it be acceptable to get the gene of interest synthesized commercially and then use it as a template for PCR or directly digest it and clone it?
Can anybody suggest how successful commercial oligosynthesis approachs this? Can I get two genes of ~ 1kb and 1.8 kb from this. Does anybody know any reliable and popular companies for this?
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In my lab we got great gene synthesis services from GENEWIZ http://www.genewiz.com/
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28 answers
control) with sequencing primers and it gave me negative results. For ligation, I use sticky-end PCR
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This ia a very common problem, when your DNA concentration is low (which is often the case after a gel extraction). The reason for this is that you have to add a large amount (Vol) of your insert (and vector). After a gel extrcation, the DNA is not as pure as you might think-in fact it is quite salty and this salt intereferres with the ligation buffer and the ligase wont work that well (they are bitches anyway :-).
But if you have many colonies for your only vector control ligation, it seems your vector is not cut well or re-ligates. Always cut over night and add Alk. phospatase in the morning for like 4h or so (directly to the restriction reaction, ignore the buffer for the Alk. phos,) then run it on the gel and extract the vector from the gel.
You should have only very few colonies for your control reaction now, but this will not solve your problem, for the ligation you have several options to improve, best to do all of them:
1, increase your DNA concentration, vector and insert (e.g. grow 3x as big cultures, run 3 PCRs or digests and pool together in a small volume) then you have to add only a little bit to the ligation reaction.
2. Don't trust a nanodrop, when your concentration is <50ng/ml. better load a few microliters to a gel and compare the band intensity with the marker (which should have a known amount/concentration)
2. Once you have a high concentration of DNA, you might want to desalt both the vector and the insert before adding it to the ligation, which will purify it even more. I use sephadex for desalting but there are many options.
3.. Always do a 20ul ligation reaction (or bigger) even if you don’t need it, because this will dilute the contamination from your DNA as well (insert and vector).
4. Ligate overnight-I normally do it at RT, 16°C, 8°C and 4°C never worked for me…but it depends I guess.
5. desalt your ligation before transformation (if you don’t and your insert DNA is contaminated with salts this will also cause the transformation to work better in the control ligation than in the actual cloning ligation)
Cheers!
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I'm trying to clean up an antibody from a serum sample. For this I have the Melon Gel Purification Kit. Has anyone ever used this one?
So I worked my way through the protocol given by the kit.
2 things I was wondering about:
1.) When I put the gel on the column, I used a 1000µL tip and it looked like it was fine. In the manual they say use a "wide bore or cut tip". Is this point important for the efficiancy?
2.) I put the gel on the column and spin it for 1 min. But the gel comes out of the centrifugation-step in a sloped way, because of the centrifugation of course. Doesn't this make the filter-effect of the gel inefficient? Like the one side is very thick and filters a lot of the serum-parts and the other part of the gel is very thin and filters less serum parts?
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Thanks a lot for the advices! I will try to focus on these next time.
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I Read several papers recently and they all used the tRNA in the hybridization buffer. Is there someone who can tell me the reason?
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Hi Xiaohui,
The tRNA is there as a nonspecific blocker and carrier to protect the probe and target. Since these are very unlikely to bind specifically to any gene of interest and were readily available and cheap from the first development of hybridization techniques this has become a standard component of nearly all such buffers.
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20 answers
Clonase reaction). I sequenced the plasmid and results from sequencing were positive for my gene. I went
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I would think that your plasmid does not have GFP. Sequence with a primer specific for the GFP codon, if you get a sequence it is there, if not you are workong with another vector than you asssume. Also perform some digests of your plasmid. Are you also not looking at endogenous expression of your goi?
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I want to estimate plant total genome size by flow cytometry. Are there any reliable, simple protocols to estimate 2C content?
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I know one laboratory for measuring DNA content in Mexico City. Is the Citogenetic Laboratory at Instituto de Biología, Universidad Nacional Autónoma de México. Contact to Dra. Guadalupe Palomino. Regards
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My set-up is mechanical stimulation of cells growing on a Petri dish. I'd like to do FISH after treatment by a centromeric probe (Cambio).
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Could you also grow your cells on slides or coverslips placed in a Petri dish?
In either case, for hypotonic treatment and fixation you have to carefully soak off the medium and slowly add hypotonic solution; after reincubation at 37°C again carefully soak off the hypotonic solution and add fixative (methanol/acetic acid). It might be best to directly add some fixative to the hypotonic solution, replacing it step by step with cold fixative, otherwise your cells might detach from the surface. Fix the cells in the fixative in the Petri dish at -20°C, then remove the fixative and air dry the cells.
Then I would employ a standard FISH protocol; however, I am not familiar with Cambio probes. I would select the area you are interested in and just add the FISH solutions to this area and use coverslips as you would do for a conventional FISH on slides. I think you have to make sure that your Petri dishes have no autofluorescence, may be you have to use a brand suitable for life imaging, may be glass bottom dishes, however, I am not sure whether this will indeed be necessary.
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I am currently going through my options for knockdown and overexpression of genes in cell culture. I am interested in the shRNA option but have some questions as have no experience of using this method.
Is there a commercial company from which you can buy ready to use (ie add to your cells) shRNA?
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shRNAs come in different types of vectors. I tried many types and got the best results with the pLKO.1 backbone also called TRC (TRC1 or TRC2). This type of shRNA vector is sold by Sigma and Open Biosystems. Usually if you order more than 5 you get a good discount. Also, if you can afford it, order more than one control shRNA since even if they should not have any effect on your cells, some of them still do. If you are not in a rush, I would not get the transduction-ready viral particles because not only they are more expensive but more importantly you may need to optimize the amount of viral particles you apply to your cells and also the incubation time. And for this optimization step, you may not have enough with what they send you. For example, too many viral particles incubated with your cells may kill your cells. Too little may not be enough to infect all your cells. To optimize the conditions of your knock-downs, it is better to prepare your own viral particles with the vectors containing the sRNAs these different companies provide. Getting the bacterial glycerol stock is usually cheaper. Then you prepare DNA from these bacteria, transfect HEK 293T cells (with the packaging vectors as well) and infect your cells with the 293T medium (different amounts, times...). If you need a good protocol, let me know.
Can anyone recommend a favourite Enhanced ChemiLuminescence HRP Substrate for nitrocellulose membrane blotting?
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I'm considering repurchasing my Peroxide & Luminol Enhancer Solutions at the moment and I was wondering what people's experiences were of using the various offerings Promega, Life, Pierce etc. have to offer. Specifically, I'm detecting HRP antibodies on a Biodyne B Nylon membrane.
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Hi Robert, Our lab uses the Amersham ECL Prime reagent (From GE Healthcare). It works for nitrocellulose membranes and PVDF ones. The ECL prime has enhanced sensitivity I believe. I can tell you that it is really strong, and often we have to dilute the reagent 1:3 or 1:4 in water to not get a glowing membrane! Because of this, we can use much less developer, and so we order it much less. Also, this allows us to either use less primary/secondary antibodies, load less protein on the gel. Plus, the ECL prime is more stable than older versions. Haven't tried many other ECL reagents, but this one works well for us! Val
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Is it necessary to generate standard curve together with samples in each qPCR analysis? The qPCR analysis is for relative quantification of a targetted bacteria population.
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Isolate high concentration Bacterial Genomic DNA (>1 ug/ ul) and dilute it in 1:0, 1:10, 1100, 1:1000, 1:10,000 and 1:100,000. Measure OD and concentration of each one DNA dilution in spectro (avoid pipetting errors ). calculate DNA copy number regarding DNA concentrations or ODs (Use online calculator http://www.uri.edu/research/gsc/resources/cndna.html) prepare SYBRGreen qPCR reaction for bacterial rRNA gene or other genes. Run PCR in 40 cycle. Set a threshold on the quantification plate. Note CT value for each one sample (DNA dilution), Set a standard curve with X(horizontal): CT and Y(vertical): DNA dilution factor.
I recommend PCR mix per reaction as: 2X SYBR Green master mix (Thermo) :10 ul, Forward primer (10 pmol/ul): 0.5 ul, Reverse primer (10 pmol/ul) 0.5 ul, DNA : 1 ug, dH20: top to 20ul. Run as 95/10 m|95/30s-62/60s-72/30s (40 cycle)
I hope this was useful
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6 answers
Rt-pcr or qrt-pcr
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The virulence of the ND virus is dependent on cleavage of the fusion site and is characterised by different sequences in the genome. Thus a reliable and rapid method to determine virulence is to sequence the fusion site. The alternative
method is to do a conventional mean death time (MDT) study, which is time consuming.
You can use rt-PCR and sequencing along with various in vivo virus pathogenicity tests. You will find many publications for discriminating between lentogenic (low virulence) and velogenic (high virulence) Newcastle disease viruses. Just google the term "F0 cleavage site and NDV virulence", you will find plenty of info!
Good Luck!!
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What reagent can replace isoamyl alcohol in DNA extraction, when used in combination with chloroform?
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3-Methylbutanol
ACS Reagent
Product Number 32,002-1
Store at Room Temperature
Exact replacement for Product Number I 0640
Product Description
Molecular Formula: C5
H12O
Molecular Weight: 88.15
CAS Number: 123-51-3
Synonyms: isoamyl alcohol, isopentyl alcohol,
3-methyl-1-butanol
This product is designated as ACS Reagent grade and
meets the specifications of the American Chemical
Society (ACS) for reagent chemicals.
3-Methyl-1-butanol, commonly called isoamyl alcohol,
is routinely used in molecular biology, notably in the
purification of DNA. It is widely used in conjunction
with phenol and chloroform, for the removal of proteins
from the nucleic acid solutions by extraction. The
addition of chloroform and isoamyl alcohol in the
extraction protocol deals with two issues that the use
of phenol alone does not completely address:
1. Isoamyl alcohol helps to inhibit RNase activity,
which phenol does not completely inhibit.
2. Isoamyl alcohol helps to prevent the solubilization
in the phenol phase of long RNA molecules with
long poly(A) portions.
In addition, isoamyl alcohol reduces foaming during
the extraction process. Isoamyl alcohol may also be
used in the extraction of ethidium bromide from DNA
solutions and in the radiolabeling of RNA transcripts in
nuclei that have been isolated from tissue.1
Isoamyl alcohol is often used in the HPLC analysis of
various pharmaceuticals and metabolites.2,3,4 It has
been used in the analysis of oxidized and reduced
pyridine nucleotides and adenylates in organic phenol
extracts from mitochondria.5
A phenol:chloroform:
isoamyl alcohol procedure for the extraction of
chloroplast DNA that avoids density gradient
differential centrifugation has been published.6
A
protocol for the isolation of mRNA from a thermophilic
cyanobacterium that incorporates a
phenol:chloroform:isoamyl alcohol mixture has been
described.7
Precautions and Disclaimer
For Laboratory Use Only. Not for drug, household or
other uses.
Preparation Instructions
This product is miscible in ethanol [0.1 ml/ml,
10% (v/v)], yielding a clear, colorless solution.
References
1. Molecular Cloning: A Laboratory Manual, 3rd ed.,
Sambrook, J. and Russell, D.W., CSHL Press
(Cold Spring Harbor, NY: 2001), pp. 1.73,
6.24-6.27, 17.27-17.28, A1.23, A8.10.
2. van de Merbel, N. C., et al., Validated liquid
chromatographic method for the determination of
bexarotene in human plasma. J. Chromatogr. B
Analyt. Technol. Biomed. Life Sci., 775(2),
189-195 (2002).
3. Dawson, M., et al., A rapid and sensitive highperformance liquid chromatography-electrospray
ionization-triple quadrupole mass spectrometry
method for the quantitation of oxycodone in
human plasma. J. Chromatogr. Sci., 40(1), 40-44
(2002).
4. De Baere, S. M., et al., Identification and
quantitation of despropionyl-bezitramide in
postmortem samples by liquid chromatography
coupled to electrospray ionization tandem mass
spectrometry. Anal. Chem., 71(14), 2908-2914
(1999). 5. Noack, H., et al., Evaluation of a procedure for the
simultaneous determination of oxidized and
reduced pyridine nucleotides and adenylates in
organic phenol extracts from mitochondria. Anal.
Biochem., 202(1), 162-165 (1992).
6. Mariac, C., et al., Chloroplast DNA extraction from
herbaceous and woody plants for direct restriction
fragment length polymorphism analysis.
Biotechniques, 28(1), 110-113 (2000).
7. Luo, X. Z., et al., Isolation of full-length RNA from
a thermophilic cyanobacterium. Biotechniques,
23(5), 904-906, 908, 910 (1997).
GCY/NSB 11/03
Sigma brand products are sold through Sigma-Ald
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8 answers
We have Corbett's RT PCR in our lab but I am having difficulty in analysis of data and I am getting amplification in negative control. Can anyone help me ?
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I use to work with the Corbett rotor gene 6000 and quick tips to analyse your data in excel:
-First in the main window (log graph) verify that you have a good amplification, the s shape curve. Adjust the threshold to make sure you are using the data within the linear range.
-Second check your melt curve to make sure you have only one amplicon. You should have only one pick if the primers are good, binding only one location. If not you data is not reliable.
Then copy and paste your data in excel and then start analysis.
-check your standard curve and make sure it is ok (R-value)
-obtain the standard curve equation and derive the concentration of your gene of interest in each sample.
-Normalize your data using an internal control (housekeeping gene) in each sample.
And that is it. But like other suggested you have to follow the manual it will tell you exactly how to do it.
Good luck
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I have included the cells in culture, in epoxy resin directly on multiwell plates. To detach, I tried using liquid nitrogen, but the plate was not broken. Do you know a method to separate the inclusions from the plate? Alternatively can you suggest a good method to include adherent cell cultures with epoxy resin without, however, bringing them into suspension so that I can observe them by TEM?
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We're trying to silence a telomerase component in HeLa cells with Lipofectamin RNAiMax and siRNA, but we haven't achieved more than 50% of silencing. We're interested in silencing about 80-90%, could somebody help us?
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hTERT silencing depends on the constitutive level of hTERT in your cell line. If your cell line contains high level oh hTERT it will be more difficult. Depending on the question you ask you have to choose a cell line containing a low expression of hTERT. However you can check telomerase activity to see whether 50% of hTERT depletion have a consequence in terms of activity.
You have also the solution to test other si RNA.
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(I need it for making competent cells)
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Autoclaving will change the pH of your buffer.. maintain sterility and try using sterile components.
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I have been genotyping flox mouse for six months. I used to get clear two bands for heterozygosity and one band for homozygous and wild type. Recently, I have started getting smears in most of the samples in such a way that I could not differentiate them. Heterozogous has 2kb and 3kb band. Interestingly, the negative control also have smears. All these smears originate from the well itself. I have changed all reagents, even tried different methods of genomic DNA isolation protocols. Can anyone please help me with this?
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Once I have that problem, somebody changed the PCR profile in the thermocycler and I did not notice it, but in other cases something as simple as change to DNAse free water works, I'd start preparing new work solution for dNTPs and primers and from that experience I always double check the PCR profile before start a new one. Good luck!
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Can anyone help me with the NaPshot Multiplex System for SNP genotyping, my question is how big should my fragment be to identify 10 polymorphisms?
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Hola Cay. Te contesto en español porque no creo haber entendido del todo bien la pregunta y no voy a aportar nada ahora. Me imagino que te refieres a SNaPshot, porque NaPshot es una siestecita. Creo que tienes o quieres 10 polimorfismos en un único fragmento. ¿Es así? Sería más normal tener o querer 10 polimorfismos en 10 fragmentos. Sean 10 polimorfismos en un único fragmento o en diez, da igual el tamaño o los tamaños de amplificación siempre que no sea exageradamente grande (miles de pares de bases) o trabajes con DNA degradado. Tampoco tengo claro si aclaro algo o me he liado por no entender bien...
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I diluted the DNA templates 10 fold until the smallest one has 1 copy and 5 copies. Then I do q-PCR with following system:
SYBR 5ul
F primer 0.4ul
Rprimer 0.4ul
cDNA 1ul
H2O 3.2
40cycles
The results are strange. The water has an obvious curve similar to 20 copies. The whole process that I did is in the super clean branch.
Can someone tell me how to avoid the negative sample pollution?
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Hi, agreed with Ajay. Take care of using ultra clean water and irradiate with UV all that you can (pipettes, tips...) also separate your working areas (obviously do not prepare your mix where you did your DNA preparations etc...).
Another clue maybe due to primers dimers. Depending on your PCR machine maybe you can do a melt curve analysis at the end of the normal cycles to check for contaminations. Or you can try to dilute your primers and check with a normal PCR if you see contamination in the negative control.
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We are beginning to work with a group of students to develop a database for us to track our samples through our very complicated analysis process. We are thinking about using Microsoft SQL server. Do you have any advice on database construction/management/usage? I have no experience with databases, so I appreciate any advice you can give me.
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Hi Elizabeth,
I would suggest that in a first step you have to define all lab processes which are supposed to be tracked in your new database. This will probably be quite a lot of work (depending on the size of the lab) as you might have to talk to a few different people to ensure that you fully understand what everyone is doing in their particular workflow. This information will then form the basis to generate a high level flow chart depicting how a sample will be handled throughout its processing (all the different possible assays, experiments, sending to other providers), as well as what kind of data it will produce and where this data ends up at the moment. The chart will probably be very complex, but it will achieve several things:
1) You will have a much clearer picture of what kind of processes and dataflows are happening in the lab
2) this will form the basis for all further discussions and planning of the software implementation
3) it will also form the basis of what exactly needs to be implemented as a minimum requirement in order to have project milestones
4) it will allow the generation of test cases to later test the functionality of the database
5) the IT guys love flowcharts and it will make it much easier for them to design a database which can handle all the processes
These steps should come well ahead before making a decision which kind of DB system or web app environment should be used to implement the solution. All the underlying logic needs to be developed before anyone starts producing code.
However, before starting to develop some custom database and frontend I would also suggest to look at what is currently available and if existing solutions could be customized to your requirements. E.g. http://www.labkey.com/ is a free solution for managing biomedical data (both research & diagnostic) but it is not trivial to set up and customize - on the plus side it might save a lot of work reinventing things that are already out there.
Hope that helps!
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I will start working with molecular phylogenetics of microalgae in a lab that doesn't have Nanodrop or any fancy DNA estimating equipment or kits. I will use a DNA multi-extractor. I previously worked with Nanodrop and the gDNA estimation was important to prepare the PCR reaction. Since I have sets of different strains (at different cell concentrations when DNA was extracted) I expect different amounts of DNA as well.
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here is what I mean. This is a pic I made in the mid1980s, i.e. in the Pre-PCR era. In this example the lowest dots show diluted DNA (mixed with Ethidium bromide) which can be compared with all the other dots above...
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I am analyzing exome sequencing data (~100x coverage). Pipeline includes BWA for alignment
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I suggest posting this question on the GATK forum <http://gatkforums.broadinstitute.org/>. They tend to be quite fast at replying to questions about their software.
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I have done phenol extraction and ethanol precipitation of a DNA sample. I have run the DNA on agarose 1% gel and excised the desired band. The band showed good intensity. The excised band was melted in in water and 3M NaAC. Phenol was then extracted, ethanol precipitated and mini-column G50 Sephadex purified. However, when I ran the DNA sample of the desired band after purification, I found no signal. It was shocking to lose such a precious sample. Therefore, I would like to ask whether anybody has any idea as how to avoid such a problem in the future?
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Add CTAB/NaCl solution to mixture. Then mix thoroughly and incubate 10 min at 65 oC. After that, add an equal volume of chloroform/isoamyl alcohol (24:1) mix thoroughly, and spin at 13,000 rpm for 5 min in microcentrifuge. Then you have to precipitate phenol/chloroform/isoamyl alcohol (25:24:1) and mix well but very gently to avoid shearing the DNA by inverting the tube until the phases are completely mixed. Also add 0.6 volumes of isopropanol to precipitate nucleic acids. Finally wash DNA with 70 % ethanol to remove excess salt and CTAB from the pellet. I'm sure your purified DNA would seen perfectly in gel.
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When I precipitate the bacteria for making the competent cells, the bacterial pellets are somewhat black. I use LB medium(10+10+5) & DH5-alpha for precipitation. I centrifuge them at 8000 rpm for 10 minutes. I'm sure that they aren't contaminated because when I use them for transformation, they work properly.
Does anyone know what's the reason for this?
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Dear Mohammad,
The brownish - black pellet you are seeing are dead bacteria.
Best regards.
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I am wondering if I mix two sirna and add to lipofectamine or seperately transfect with other siRNA the next day.
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I guess yes you can if you're using single siRNA (not pool) where you use [5nM] of each type and then transfect using lipofectamine 2000. You can always try Dharmafect 3, it has a good transfection efficiency (sometimes better than lipofectamine 2000). However, I don't recommend that you transfect with one siRNA and the second day you transfect with the other one because you'll be stressing the cells and cells may even die. Additionally, try to do reverse transfection instead of regular transfection (add lipid/siRNA in the plate and then add the cell suspension on the top - time of transfection is really depends on what you're looking for - in my case i do 72 hours).
Hope that helps :).
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I design two primers with two RE site in 5' ends
I've used a double digest reaction for cutting my PCR product and after running on 1% Agarose gel
I used it in ligate reaction as a insert fragment
How can I be sure that pcr product has been cutted properly with both of my RE?
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Yes it is possible. If the RE leaves 5’ overhangs you can fill-in with a labelled dNTP in the dNTP mixture and incubating with Klenow in the appropriate buffer. It depends on the method of detection facility you have access to. A Taq polymerase which doesn’t add an extra terminal nucleotide can also be used. You should not add primers in this case.
If the RE leaves 3’ overhangs, it is a bit laborious but doable. Alternatively if you only cut from one end and your fragment is not very big you can separate them on a denaturing urea PAGE or a sequencing gel.
As you see it is worse the remedy than the illness and it would be absurd and nonsense to perform such experiments only to clone a fragment. Instead, I suggest that you follow some suggestions above. What you are doing has been done hundreds of thousands times and it worked for most people. So, check your RE on a plasmid to make sure the enzyme has not been left out the freezer, etc...Also on the enzyme catalogue you should respect the manufacturer instructions relating to the digestion at the end of a DNA fragment, star activity, etc... Some enzymes may need more extra nucleotides upstream their restriction site than others, etc.... Try to purify your fragment on a spin column (Kit) after each digestion. Normally 1 hour should be OK.
To make sure your PCR product is OK, I would follow Arjan’s recommendation. You can sequence first your fragment and then cutting it from a plasmid will circumvent your issue..
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There are many formulas and program applications to compute melting temperature and annealing temperature of PCR primers nowadays, but these tools usually have different results. Would you be so kind to give me some information about how to calculate this values correctly? These temperatures depend only on the sequence of primer or on other parameters too?
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Hi! Andrii,
There are many software's, if you Google you get hell lot of information. Do not worry about that.
So far what ever the primers I designed are using by Primer 3 (NCBI).
Submit the sequence, it automatically calculate Tm, do not worry how to calculate for that (you had standard formulas for all), just reduce 5 degrees to Tm then you get Ta and set PCR it works very fine.
I designed many primers, they all worked very fine, got very good results.
Try it. You can save lot of time. I am sure you get nice results.
Good Luck!!
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I normally inoculate 5 - 7 colonies from selective antibiotic LB plate after transformation in the liquid LB with the same concentration of the same antibiotic. The next day, I do plasmid miniprep for plasmid isolation and then run a restriction digest for all of them and go for the pattern that is expected (from VECTOR NTI). If found, I sequence and confirm it, then save it in the stock with the proper numbering and labeling.
Later, I pipette 1 ml of the culture from the above liquid LB media of the same plasmid that has shown me the expected banding pattern and inoculate again in 130ml of LB and the same antibiotic (and the same concentration). I do plasmid midiprep and isolate a high concentration of the Plasmid.
Out of curiosity, I run the undigested and restriction digested miniprep and midiprep to confirm. Here my undigested shows the similar results but my redigestion of midiprep varies from the miniprep and the expectation. Any comments on this?
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Your DNA is correct ! It has not been digested properly. In fact, if you have the same gel, increase the exposure time and capture the image; you will see the desired bands. I could see it by adjusting the contrast of the image. There is still lot of supercoiled DNA left in your gel. You can dilute the DNA and re-digest it and you will see the desired product.
For your reference, I have attached the edited gel picture. Hope this helps.
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I am using a thermocycler from BioRad (USA) and PCR tubes from TARSONS (0.2 ml volume). Reaction volume was 25µL, please suggest some solutions.
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lid temperature must be high than plate
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I want to clone a 300 nucleotide fragment in pHannibal vector with xhoi and EcoRI restriction sites. I designed two primers with these restriction sites in 5' ends of my primers
Forward:CGGGCCGGATCCCTCGAGG......
Reverse:TTCAAAGCTTGAATTCCAT....
I cut my PCR product with these enzymes with fermentase recommendation protocol. For double digest: 2 x tungo + 1u enzyme for five hours at 37°C. Also, I cut my vector with these enzymes separately, firstly with XhoI and after that with EcoRI. For double digest: 2x tungo+1 uXhoi==>20 reaction for fifteen hours ==> inactive with heat 80°C for fifteen minutes >> +1u EcoRI for fifteen hours. After that, I extract cut the plasmid and my digested PCR product from gel and ligate them. With fermentase t4 ligase according to the manufacturing protocol: 1 u t4 ligase+ 1x buffer+(1:3) ratio. Finally, transformation with ligate product had a lot of self ligate colonies. My attempt to use another ligate ratio (1:1) or with high insert has not any colony with inserted. Also, my attempt to clone my fragment in pblusescript sk+ has similar results. I had a lot of blue colonies in my plate, what is the problem?
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Hi Mohammad,
sometimes, there can be problems, if the two restriction sites are located very close to each other. Some enzymes require a restriction site to be located a certain distance from the end of the fragment (general rule 6bp), otherwise they cut with a low efficiency. you might try to read something about troubleshooting:
Recognition site may be too close to the end of the DNA fragment (for example this: http://www.neb.com/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments).
From my experience once i had a similar problem, when EcoRI did not cut after digestion with another enzyme. You could try to cut your vector first with EcoRI and then with XhoI.
Hope it'll help, good luck.
Andrea
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I would like to overexpress a gene in cell culture model. I have never done this before and from what i have read so far it typically involves establishing a stably expressing cell line usually via retroviral methods. I currently dont have the experience to do such an experiment and so was wondering are there any commerical options available to me whereby i could give the sequence of the gene to the company and they generate the cell line for me. Or if there are other suggestions about what may be the easiest straightforward method for me to achieve my goal would be appreciated.
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well, you dont have to estabilish a stably expressing cell line, if you want to see the behaviour of your over expressed protein inside the cell line. For transient expression of your protein (human?) you can simply clone it in pcDNA vector(driven by CMV promoter), and transfect them in a mammalian cell line (eg:HEK 293T), in one or two nights, lyse the cells and do your assays with the protein in the cell lysate. It all depends on what you want to do further. Expression of two or more proteins with the respective vectors can also be done using this.
Does anybody know a good enzyme for high-fidelity long-range PCR?
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I need to amplify a 7kb cDNA from a plasmid in order to sub-clone it into another plasmid. I searched in google and found some options, but I'd like to know if anyone has experience with long range PCR and could recommend me an enzyme that works well for the task (amplifying 7kb with few or no errors).
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I've been using Phusion High Fidelity DNA polymerase. I've had consistent results using this enzyme. http://www.neb.com/products/M0530-Phusion-High-Fidelity-DNA-Polymerase ;-)
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Dictyostellium is an soil living amoeba and the basic use of this organism is DNA repair, signal transduction, transcription etc. I know the GST-pull down method, but through this technique the antibodies bind with partners instead of direct attachment.
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hello:
You also can detect the protein protein interaction by Far western. It can be a direct way.
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I am currently working to optimize an indirect ELISA in order to detect the amount of specific IgYs present in enriched serum from eggs of chicken inoculated with synthetic peptides.
Do any of you have recommendations for the optimization of the assays? Particularly, concerning the concentrations of the peptide for coating and the buffers that it is better to use. I am having trouble with the coating of the peptides to the plate. For some of them, coating with normal PBS works fine, but for others, I don't get any signal at all.
Also, when I do a standard curve with IgYs only (0 to 60ng per well), the signal is too strong and I can't detect any difference above 10ng.
I'm using high-binding Microlon plates, PBS 1X (10mM phophates) for coating, PBS only for washing, PBS with tween-20 and gelatin for blocking, and Donkey-anti IgY with HRP in a 1:2000 dilution as a conjugate.
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Hi Johana,
One problem may be that some of your peptides are too short or not hydrophobic enough. We routinely coat microtiter wells with peptides that are 15-25 aa long. Another buffer to try diluting your peptide in is a 0.1 M sodium bicarbonate buffer, pH 8-9. The ELISA plates we typically use are Immulon 2HB or 1B, depending on the assay. Here is a quick reference guide to the differences between the types of plates we use and their properties: http://www.nuncbrand.com/en/frame.aspx?ID=12001 . Not all plates are the same and the type of plate can greatly affect your results. Plate binding properties can also vary from manufacturer to manufacturer. So another factor may be that the high binding plate you are using is contributing to the high background binding with your IgY's.
We've had to test a number of different ELISA plates to get the optimal adsorption of peptide and low background. It would be worthwhile to try out plates with different binding properties.
When coating peptides we usually make a 100 ug/ml solution in sodium bicarb (or PBS for some peptides that we use) and add 50 uL/well to give a final peptide concentration of 5 ug/well. Coating is done overnight at room temperature. One wash with your coating buffer should be sufficient before blocking. We make up a 3% BSA blocking solution in the same coating buffer that the peptide was made in. In between incubation steps, we wash 3x by adding 200 uL/well of TBS or PBS containing 0.1-0.05% Tween-20 (you can adjust the amount of tween to see if it helps with the high background.) Hope some of these suggestions help.
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If I wanted to create an auxotrophic mutant like dapD , do I need to add the isomer downstream from the product in order for it to survive? Do I need to add all isomers or just one? I'm not sure which compound I need to add in the medium to disrupt the dapD gene, so I've attached the link of the pathway. Does anyone have any suggestions?
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You should be able to add any one of the intermediates that is on the pathway for your organism after the knockout. So L-L-DAP or meso-DAP should both work in virtually any organism.
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Is it possible to use as probe a PCR fragment obtained with a primer 5' labelled with CY-5?
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Hi Violaine, sequencers are essentially fluorescent imagers with a one-dimensional scanning area. Chromatograms are a plot of retention time for a nested set of fluorescent polymerase products. If the protocol works on a sequencer, it should work on a fluorescent scanner. If you scan with an above-the-surface offset, you might not even have to remove the gel from the glass plates.
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I want to transfect THP-1 cells (and if preliminary experiments work, I plan to later transduce them with a virus to generate a stable cell line). For my functional experiments, I want them to be differentiated into adherent macrophages using PMA.
Ideally, I would like to transfect the adherent cells, for sakes of the time-course of knock-down.
I understand that THP-1 suspension cells are very difficult to transfect using lipid-based reagents, but nucleofection works well. I would like to know if once they are PMA-treated, whether the cells will be more amenable for transfection with lipid-based reagents. I don't really want to differentiate the cells, get them back into suspension for nucleofection (trypsin or EDTA?), and then re-plate. Any suggestions?
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I have just tried to electroporate THP1 and induce the differentiation right away after electroporation. According to the manufacturing procedure, this should work. I will let you know if this works well from my side.
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We are trying to quantify the amount of nitric oxide produced by our cells following an inflammatory stimulus. To do this we used the modified Griess reagent from Sigma. Upon incubation with the reagent for the suggested 15 minutes there was no colour change in any of our treated cells. However upon the addition of more reagent or upon incubation for much longer (an hour) there were colour changes indicating reasonable levels of nitric oxide. In all cases the standard curve did not change colour (after the first fifteen minutes or upon further addition of the reagent). What should I make of this- is this a meaningless quirk of the reagent or could it represent a real signal (perhaps from a nitrate or chemically similar precursor)? I would have dropped this but we believe from the literature that NO is induced by our cytokine stimulus in our human hippocampal stem cells. Any advice would be great.
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Mark
The good news is that you only get the purple color from nitrite by diazotization, so if you are getting purple color it is coming from nitrite and not some unrelated compound.
Have you checked pH? The reaction needs acidic conditions and the symptoms you describe suggest your media may be buffering the acid and causing a slower reaction (fixed when you dump in more reagent).
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I am looking for a kit to do linear RNA amplification. Specifically I am trying to decide between the Life Tech Arcturus RiboAmp plus kit, the Qiagen Quantitect whole transcriptome kit, or the Clontech mRNA amplification kit. I have also heard Ambion makes a similar product. Does anyone have any experience or recommendations?
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We do our amplification using the arcturus kit. It gives verry good results and we can start we verry low RNA quantity. We ensure the RNA quality of the RNA doing a Bioanalyzer first to the amplification. Just be sure before choosing a kit that you want oligodT priming or random.
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After thermocycling there were IC bands in all but not in the NC, I repeated the procedure and the result was the same. Is there any kind of inhibition in Mastermix?
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If you have probably included the IC during extraction of nucliec acid for all samples and have forgotten to include your NC during extraction, it could be the reason for absence of IC in your NC !! Since you might have directly taken water or Negative template control for PCR directly rather than including that for nucleic acid extraction step (which includes addition of IC). Could you be more clear in what protocol you followed i.e.when you use the IC ? That way we could help you better.
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Have been using the semi-quan API-ZYM test to see the abundantly expressed enzymes by the microbial consortium in a lab-scale Rc treating wastewater. The enzyme "Naphthol-AS-BI-phosphohydrolase" keeps popping out as one of the abundant enzymes, no matter if the feed is synthetic or real wastewater. I cannot find solid info on what this enzyme does. Any idea about that? PS: the substrate of the enzyme in the API-ZYM slot (no12) is "Naphthol-AS-BI-phosphate"
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Hello I am also in the same situation ultilizando sludge on soils with different sludge treatments that enzyme being of the highest active hope can also help us with more information about it. Cheers
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I tried to research the halflife of a protein. This surface protein is expressed continuously by cells. So an idea is the biotinylation of the protein in cell culture. I am absolutely new in this field. So maybe you can help me understand how this works.
1. How can I biotinylate only the target gene?
2. How can I measure the halflife?
3. Any literature or protcols which are good for persons who are new in this field?
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Cell surface biotinylation assays: Cells were washed three times in ice-cold HBSS and then exposed to 0.5 mg/ml Sulfo-NHS-SS-biotin (Pierce) for 30 minutes at 4 °C. To quench excess Sulfo-NHS-SS-biotin, cells were washed three times in HBSS with 5 mM Tris (pH 7.4). Following this step, the procedure was adapted to the goal of the experiment. (1) For determination of cell surface receptors, cells were lyzed in RIPA buffer and centrifuged for 20 min at 20.000 x g. Clear supernatants were incubated with EZview Red Streptavidin Affinity Gel (Sigma) for 4 hrs at 4°C. Beads were sedimented by centrifugation at 1000 g for 5 min, and washed four times with RIPA buffer. Aliquots of input and precipitate samples were analyzed by Western blot using appropriate antibodies. Chemiluminescent Western Blot signals were quantified by a ChemiDoc XRS luminescence imager (BioRad) in combination with Quantity One software. (2) for endocytosis: Cells were stimulated with agonist in prewarmed (37 °C) DMEM growth media for the appropriate time at 37 °C. Remaining biotin label from cell surface proteins was removed by washing the cells twice in ice-cold glutathione stripping solution (50 mM glutathione, 75 mM NaCl, 1 mM EDTA, 10 % FBS, 75 mM NaOH; 2 x for 30 min at 4 °C). To remove residual glutathione solution cells were washed twice in ice cold HBSS. Cell lysis and quantitation of biotin-labeling was performed as described in 1. (3) for degradation: Cells were stimulated with agonist as above. Remaining biotin label from cell surface proteins was removed by washing the cells twice in ice-cold glutathione stripping solution (30 min at 4 °C). Glutathione-stripped cells were again incubated with prewarmed growth media for 3 h or 6 h in a CO2 incubator at 37 °C, followed by cell lysis and quantitation of biotin-labeling. Ref: Turvy DN, Blum JS. Biotin labeling and quantitation of cell-surface proteins. Current protocols in immunology / edited by John E Coligan [et al 2001;Chapter 18:Unit 18 17
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I am getting a background smear besides specifc bands when I run DGGE gels. The PCR products used for DGGE gels were obtained by regular PCR using Taq pol with GC-clamp primers from genomic DNA isolated from mixed species biofilms. DMSO was used. Any suggestions?
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You could also try to add a 80 degree step in your PCR (after elongation) this could help to reduce primer dimer formation, I did that in G. Muyzer's lab. For your PCR: Do you use Primer for 16S? Your DNA is from biofilms? So the smear could also be an indication that you just have a really high diversity. But try to increase your annealing temperature, maybe add the 80 degree step, load different amounts of DNA on the gel and then have a look ;o) Good luck!
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I am working on Burkholderia, endosymbiont in the subgenus Crispardisia. I would like to know if it is possible to track the circuit of the bacteria in the plant as it is impossible yet at this stage to isolate it. The bacteria already are in the seeds and propagate while the plant is growing, and will settle in the leaf margins, apical shoot, and axilary bud. So I would like to understand how the bacteria propagate, without having to isolate it first.
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One suggestion would be to use immunofluorscence -there must be some property of the bacterium that you could use to detect it. An antibody that crossreacts? A Lectin that has binding to the LPS and not the plant? Even without that you should be able to visualise the bacteria in the plant tissue by microscopy. Unfortunately, any method would be destructive for the plant.
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I am planning to do a loss of heterozygosity (LOH) analysis for the CDH13 gene at different polymorphic microsatellite markers in ovarian tumor tissue. Can anyone suggest which is the better option for LOH analysis: conventional PCR or Real time PCR?
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Real-time PCR works:
Wilhelm J, Pingoud A, Hahn M. Detection of p53 allele deletions in
human cancer by quantification of genomic copy number. In:
Meuer S, Wittwer C, Nakagawara KI, eds. Rapid cycle real time
PCR: methods and applications. Berlin: Springer Verlag, 2001:
159–70.
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We are trying to enrich for very rare viral transcripts in RNA extracted from a frozen piece of tumor. Due to the low expression level of these genes relative to cellular genes, we have a hard time finding an assay sensitive enough to consistently and quantitatively measure transcript levels.
Has anyone ever tried suppression subtractive hybridization for similar problems? It seems to be a complicated assay, so I am cautious about investing in the necessary materials unless I am confident it will help. Also, in the protocols I have looked at the final subtracted cDNAs, subcloned using the attached tags - has anyone ever tried to use a biotin tag or something similar to purify these cDNAs as opposed to cloning?
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We used SSH in combination with cDNA microarrays in several studies for analysis of differential gene expression. Generally, this technique is working very well. However, if you are not able to quantify your viral transcripts using qPCR (Did you also try Taqman assays?) I would suspect that you won't be successful with SSH applied to your specific problem. You will get many false positives in your subtracted library and won't get quantitative results.
You could try Illumina RNA-Seq. Sensitivity increases with the number of reads. From one HiSeq lane you can get more than 150 Mio reads.
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There are no recommendations in the kit I use (Epicentre MasterPure). I would like to run 24 samples at once, would that be stupid? Does anyone have any experience in this?
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I prefer do that with high care to avoid cross reaction between the sample you work on it and give accurate extraction
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We're doing RLBs (Reverse Line Blots) which are a cross between a Western and a 2D blot, but using DNA oligos (probes) and bacterial PCR products (run 90 degrees to probes). We are doing a multiplex PCR using a genus-specific 23S-5S spacer primer set and a universal eubacterial primer set. We are trying to determine which bacterial species is present in a sample. We sometimes see one of 2 probes (never any others) lighting up on the blot in the no DNA lane, but NOT in the other lanes. We've tried fresh water, wiping with 70% EtOH and a Nucleoclean solution and UV irradiation. We've run out of ideas and could use some help.
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When dealing with contamination, it is best to start with all new ingredients of the PCR mix and discard all the old reagents. This includes primers, probes, dNTPs etc. If you have already done this and you still have problems, Abdelhalim may have the solution.
A likely cause of contamination when using universal eubacterial primers is amplification of E. coli DNA that is contaminating your PCR mix. The Taq in your PCR mix is generally produced from recombinant E. coli cells and some nucleic acid can be co-purified when producing the enzyme. One solution, apart from that presented above, is to purchase a PCR mix where the Taq is purified from T. aquaticus and not E. coli. Takara used to supply such an enzyme but I am not sure if they still do.
I hope this helps.
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I am interested in investigating the role of physical exercise on MMPs and it is well-known that after over exercise MMPs (mostly MMP2 and MMP9) expression increases. My interest on the cellular dysfunctions (not diseases) mediated by MMPs during/after over exercise. Which points and/or dysfunctions do I look for?
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Anyone please....
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According to MIQE guidelines, in qPCR analysis, we need to have triplicates for standards. How about the samples? We need to have triplicates for each sample as well? Can we go for duplicates?
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Hello Ying,
you got in essence the answer you were looking for above as people gave you correct answers. But for a more "educational" answer let me precise things:
1. The technical replicates. The reason one performs these replicates is to control for the validity of the method. This is the sole purpose of it. For example, having 3-plicates is somewhat of a standard. Why? Let's see: You get 3 Ct values out of your replicates: 27,3; 27,9; 35. This was pipetted from the same cDNA. So obviously, something wrong happened to the 35 Ct value. You can probably confirm it with the melting curve. And in any case, it is fair (not perfect!) to eliminate your 35 Ct value from your average. Now imagine you did a 2-plicate. Your values are 27 and 35. What is the good one? You average them? No, the only option would be to set this sample aside from analysis. You just lost a biological replicate to save on a technical replicate... It is in general far more expensive to generate a biological replicate than a technical one. Do not cut on technical replicates (that is my advice as it is not cheaper anyway). Or at least if you do, expect to have some samples that will be unusable.
Nota bene: The only valid statistical operation to do with these Tech Replicates is the average. They do not control anything else than the method's error (and pipetting, included in the method variability for me). Do not do stats analysis on it as it is meaningless (unless you try to control the machine's performance...).
2. The biological replicates: This is highly variable. According to your model, you will need at least 3 biological replicates. No statistics can be done on less than 3 samples in any case. Example, you want to compare IL-12 expression in macrophages stimulated or not with LPS. Then you need at least 3 untreated cultures of macrophages and 3 LPS-treated cultures. But very likely, you will need more for subtler effects where the gene induction is less than 10-fold. Plan 6-10 biological replicates for something that is not very strong.
As C. Thompson mentioned, you will detail how you performed your experiment in your material and methods, and you can choose to do less technical replicates, as long as you know what you risk.
As O. Berkovitz mentioned, having more biological replicates is often the key to success as this is where the most variation is observed, never neglect this neither.
On that, good luck, you can now make an informed choice and live with the variability that you choose!
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We are using the Leica Microdissection system which drops cut out cells into lids of 0.5 ml PCR tubes, that are secured in a special metal holder. Now that our batch of Eppendorf tubes have run out we wanted to re-order them but turns out they have been discontinued. The new Eppendorf 0.5 ml PCR tubes (Cat nr. 0030124537) have a bigger lid that doesn't fit in the holder. Does anyone who uses this sytem has an alternative?
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hi, we also have an LMD6500 - we use the Greiner thin wall tubes, as specified by Leica (Cat# 682 201)
josef
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I constructed bacterial cells carrying His6-tag at cell surfaces and would like to recover them out of cells that do not have His6-tag at surface. In the next step, I will isolate genomic DNA from these cells and identify insertion sites at surfaces.
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Dear Br. Suwat,
You can use Ni-NTA agarose beads.
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Is there any way of transforming your constructed DNA into the Bacillus cells other than natural competency? I have been doing natural competency using Bacillus, but I am not getting successful results.
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B. subtilis is worldwide transformed exploiting its natural competence. It would be good to know if the problem is the strain you are using (may be it is competence-deficient) or the DNA you are trying to transform. Have you tried a couple of different strains with plasmids or markers know to work in Bacillus? If you do not succeed in this way, you should look at the quality of the components of your growth medium. Finally, electroporation of B. subtilis has been tested (Biotechnology Techniques Volume 5, Number 1 (1991), 9-12, DOI: 10.1007/BF00152746).
Good luck with your experiments!
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I am working on post-transcription regulation of mRNA encoding big protein (app. 110 kDa) in Escherichia coli. I expect that upon stress conditions the ribosome employs different start codon on mRNA which is located upstream of the start codon used under normal conditions. We have analyzed ternary complex formation between the ribosome, initiator tRNA and mRNA using toeprinting and primer extension. It supports our data about using the alternative start codon. Translation initiation on the alternative upstream start codon leads to addition of 8 amino acids to protein's N-terminus. Now, we would like to distinguish between two proteins, between initial form and the form carrying additional 8 amino acid residues on N-terminus. It is desired to have a fast method, at least as fast as 1D SDS-PAGE. As our protein is too large, we are not able to distinguish between two forms using 8%, 10% or 12% Tris-Tricine SDS-PAGE. Now we would like to fuse first 10-15 codons of our protein to a small inert folded protein with MW app. 10-12 kDa. Do you have any suggestions?
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If you think GFP is to big you could use the splitted vertion of GFP (the one that is usually used for Bimolecular fluorescenc complementation experiments). I am possitive that anti GFP antibodies recognize at least one of the two halves of GFP protein. The Mw is around 12-15 KDa. Another option might be trying to clivate your protein with a protease or a chemical method. If you can clivate your protein just in a few sites you might be able to pay atention to the N-terminal fragment and see the difference. Another option might be adding an speciffic protease sensitive secuence inside your protein f you have idea were you could do that without altering its folding.
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I want to pull down a membrane bound cytoplasmic GFP tagged protein. For this I ordered GFP tagged beads from chromotech. I am lysing the cells expressing GFP tagged protein of my interest with lysis buffer (150mM NaCl, 1mM EDTA, 1% Triton X-100, 50mM Tris-HClPh 7.4). After that i incubate the protein lysate and the beads tagged with ab for GFP at 4 degree for overnight. Next day i give it 3 washing (washing buffer : 250mM NaCl, 1mM EDTA, 2% Triton X-100, 50mM Tris-HCl Ph 7.4) to get rid of unbounded protein. Finally i boil the beads for 10 min at 90 degree with 2x sample loading buffer with 4% mercaptoethanol and separate the proteins in 10 SDS PAGE. But I am unable to see a good intensity band that i can send it for MS. Even i pull down cells to increase the band intensity but there in no improvement.
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You might try the following: use 0.5% NP-40 as detergent instead of Triton X100 (it should interfere less with the ab-binding), 2nd: for the washing - reduce the NaCl concentration to 150 mM (but not below) - that is the physiological ion strength - and might improve the binding to the ab.
Good luck, Johannes
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Considering the hazardous nature of phenol, I don't want to take any risk of half knowledge. If anybody here has made the Phenol/CHCl3/Isoamyl Alcohol (25:24:1, v/v) solution by himself, please share your practical experience with me. I have phenol, chloroform and Isoamyl Alcohol separately. Now, I want to know a stepwise protocol to make a Phenol/CHCl3/Isoamyl Alcohol (25:24:1, v/v) solution. The only problem I feel is how to make the correct buffer saturated phenol for this solution. Please could anyone include all the information about what to do and what surely not to do?
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Dear Yogendra
I am old enough to give you this recepy, as I did this solution by myself - along time ago.
First of all, you need the "Chloroform (24+1)" solution which is prepared as follows:
mix 240ml Chloroform with 10ml Isoamylalkohol.
Second, use the following ingredients to make yourself the "Phenol" solution:
Use 40g Phenol (solid), 10g LowTE buffer (10mM Tris HCL pH 8.0, 1mM EDTA pH 8.0) and 50 g of your existing "Chloroform" solution.
Stir it (even by slightly heating) and then filter it into your glas bottle. We always used some staniol paper (wrapped around the bottle to protect against light) and kept in the fridge at 4˚C.
Any further question?
my best
Rolf
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the primer sequence and hence can't go for sequencing? Is there a way to find out these unknown amplified
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Probably you can clone the products into a plasmid vector and then sequence them with a single primer from the plasmid. (Obviously, you can't sequence with any primer mix intended for PCR, since it contains primers for both strands and will produce the mixed sequence of both strands and be unreadable.)
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I have been getting contamination in my PCR reactions. Because I could not eliminate the contamination internally, I bought new a pipet, new filtered tips, new SYBR, new water and new primers. However, I am still getting the unexpected amplification curves in my reactions! I am using primers (25nmole DNA Oligo) from Life; PCR water from Ambion; Power SYBR Green PCR master mix from Applied Biosystems. On the master mix bottle there is a label saying: PCR master mix (2X). Before using SYBR from Applied I used to use master mix from Qiagen. Can the new SYBR produce false positive results? If yes, why? Can anyone help?
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Everything before is right, primer dimer is classic case of amplification in negative controle, but it normally appears late in cycle number, and the melting temperature is different from target amplicon. Moreover you can check the size of the amplified product on agarose gel to see if it correspond or not to amplicon size.
On the other hand when you perform the PCR of the same target a lot of time, for long period, you should have aerosol of the target product in the PCR room. So as suggested by Anna, it is important to have different room for PCR preparation, run, and analyse. If your contamination persist, just try to prepare your PCR reaction in a never tried room, even in an office!!!
Good luck!
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I have a problem in the separation of small DNA by Agarose Gel Electrophoresis (though I optimized with different concentrations of gel). Separation was very poor and always very blurry, smear bands with DNA ladder (every 20 bp scaled up from 20 to 200 bp). I plan to run with SDS-PAGE with thought it will be better, but not sure it will work well. Another problem of using SYBR gold dyes which seems less effectively staining DNA, RNA than using EtBr which we do not want use in our lab. Highly appreciate suggestions.
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Hi, Kien. I used silver staining of nucleic acids for detection of SNPs after PCR of alleles. it worked well both for heteroduplexes and for diferenciate secondary structure in sisngle strand DNA. I know that in the "prehistoric ages" of manual sequencing it was used instead of radiactive labelling.No idea if it works with RNA. I don't know were are my notes but in "molecular cloning. A laboratory manual" by Sambrook and Russell it's described a similar protocol (Appendix 9.6):
Rinse the gel in water twice, fix the gel in 10% ethanol (10minutes), remove ethanol and cover the gel in 0.7% nitric acid (6min) remove and pour silver nitrate 0.2% (30min), remove and cover with 125ul of formaldehyde solution (37%) diluted in 100ml of developer (22.9g sodium carbonate per liter) and wait for color development.
all steps in a rocking platform or shaker. To stop the reaction remove the developer and add acetic acid 3%. Good luck
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The antibody (and exact protocol) that I'm using worked in the recent past, but now I get very high background, along with a "ladder" of negative ghost bands against the dark background. My protein of interest still faintly shows up as a positive (dark) band, despite all the non-specific ghost bands. But from my reading, ghost bands supposedly indicate the binding of too much secondary antibody. Does this imply that my primary antibody is somehow preventing binding of the secondary? By the way, the secondary is working fine with other primaries.
I've attached an image of the scanned film: lane 1=protein positive control, lane 2=knockout tissue, lane 3=wildtype tissue, lanes 3,4=experimentals.
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I recently had a similar problem. As is standard with western blotting, there are a few potential causes for the non-specific bands. Unfortunately there is no "quick fix" and it really is all down to trial and error to see what will work best for you and the antibody in question.
1. The membranes aren't sufficiently "blocked" prior to incubation. Do you include any type of blocking agent in with your antibodies? I usually include 2.5-5% non-fat milk or BSA in both my primary and secondary antibody.
2. The concentration of one or both of your antibodies is too high. If I ever suspect this, I first try diluting the secondary antibody. It might also be worth doing some dot blots to find out exactly what the optimal concentrations are for your antibody combination. You never know, the manufacturers may have changed something, or if you are using a different brand of secondary antibody that might also affect the binding properties.
3. You are not washing the membranes for long enough in between incubations and/or before exposing the blots. After my secondary incubation I usually allow the membranes to wash for a couple of hours in TBST at room temp. (changing the TBST every 10-20 min). If I still find that I have lots of non-specific bands, I will leave the membranes overnight to wash at 4 deg C. That usually cleans the blot up enough to get a clean image. If the antibody is particularly problematic, I'll also wash for an hour in between primary and secondary incubations.
Personally, I'd begin by giving the membranes a good wash and if this isn't successful, start again and this time make sure everything is blocked and washed sufficiently. If you are still having problems after that, try reducing the concentration of your secondary antibody.
Lastly, even if the antibody has worked a hundred times before AND with the exact protocol you are using, the western blot god is a cruel mistress. Maybe you're not wearing the right socks on the days it hasn't worked? ;) Good luck!
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I have obtained a few Cryptosporidium parvum MLST data from three countries by 12 microsatellite and minisatellite loci. I want to know how to achieve an existing transmission association among the three countries according to the genetic diversity data? In addition I also want to know how to estimate the splitting time of Cryptosporidium based on MLST data and the genetic data of Apicomplexa parasites.
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In relation to transmission association between the countries you can test for either genetic relatedness using Fst or Gst. However I prefer the one where you test for isolation by distance but all of them should be able to tell you whether your populations are interbreeding or not. You can use the freely available and easy to use program called GENEPOP to do these tests.
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miRNA isolation from tissue samples
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You can use standard protocol for TriReagent (TRIZOL etc.) and further isolate using BCP or chlorophorm. No columns required, but you must check the quality (DNA and protein contamination) of obtained total RNA including small RNA fraction.
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I am now looking for a generation of memory OT-1 CD8 T cells in vivo and then adoptively transferring them to B6 mice. But all the literature I read so far says to first transfer OT-1 Rag -/- cells to congenic mice and immunize the mice with a particular antigen, namely, ovalbumin expressed by a microbe such as listeria or VSV. 60 days after the primary immunization, they said enriched CD8 T cells containing OT-1 memory CD8 T cells will be used in an adoptive transfer study.
I do not know if these transferred cells are all OT-1 specific or if it is a mixed population of OT-1 specific memory CD8 T cells plus CD8 T cells from the host mice.
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I don't think you need to worry about purifying OT-1 cells, just purify CD8 cells if you want. You can separate out which are the OT-1 in the end by which bind ovalbumin tetramer or which proliferate in response to ovalbumin peptide.
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I am trying to deplete specific DNA sequences from a mixture by hybridizing with a biotinylated ssDNA fragment at 65 degrees, 180mM KCl and then capturing with streptavidin magnetic beads. The trouble is, I'm getting way too much background. The protocols for capturing biotinylated DNA with these beads typically employ 1M salt - why so high? I'd like to reduce it because I believe it's causing non-specific hybridization.
Alternatively I could raise the temperature (I used 15 minutes @ 42 degrees for the bead capture step) or add formamide, but I don't want to denature the streptavidin. Maybe I could add random hexamers or something. Has anyone tried these or come up with another solution?
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I can probably help you but need more info. So, the fragments you are trying to capture are the one that you don't need? In other words, you are cleaning the mixture from them and what is left is what are you interested in? How your your fragments in the mixture were obtained, by use of restriction enzymes or PCR? In general, the high salts increase stringency, and that should in effect reduce the non-specific hybridization.
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I trited to optimize the system by using 2500ng of plasmid DNA with different amounts of lipofectamine 2000 (5ul, 7ul, 8ul, 9ul, 10ul) in the 6-well plate. There is a light band showed up in the well when I used 8ul and 9ul of lipofectamine. However, the intensity was as low as the background. Can anyone help me to optimize the transfection system? By the way, the protein size I tried to transfer is 65 K.
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There are many things that can go wrong when tranfecting Hela cells.
I will try to answer your questions in the approximate order of the exp.
1) This might sound obvious but, double check that you have the right plasmid, and that start and stop codons are correct. Double check your sequence, do not trust that some one gave you the right thing, mistakes happen. Also ensure that the plasmid has been purified from a kit with low bacterial endotoxins in the final product
2) Ensure your cells are free from microplasma infections as this can disrupt/reduce transfection efficiency. (this can be checked either by a commercial kit pcr based kit, or methanol fix cells on a cover slip stain with DAPI and look for small speckles on the slide, that are not part of the nucleus).
3) Ensure that you have cells at ~90-95% confluency, at the time of transfection. Seeded one day prior to your exp.
4) Ensure that you seed cells in antibiotic free media
5) If you follow the printed protocol of preparing lipofectamine 2000 in serum free media like optimem, try the suggested ratios of 1:2 or 1:3 (dna µg :Lipo µl) and then keep to the incubation times they should suggest. and presuming your plasmid is correct you should get some expression
6) How are you detecting your protein? is it tagged or are you using an antibody against the endogenous protein. If it is tagged, get a positive control plasmid from someone.
If you follow these steps and still can not see a band double check your Antibody?
The only other thing could be that your protein is highly regulated and is being rapidly degraded. In this case you could add lysosomal and/or protease inhibitors to transfected cells and see if this increases your detected protein levels.
Neil
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I'd like to receive some input from people who worked with hybridomas. I need to resurrect them from being stored at -70C for about 5 years. I've never worked with hybridomas before and would appreciate any suggestions on what would be good growth factors, conditions and/or techniques.
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Should be fairly simple. Normally you would just thaw them like any other cell line and put them into RPMI1640 + 10% FBS + non-essential amino acids + 2-mercaptoethanol (for better antibody quality, not needed during simple growth) -> look for recipes ar Invitrogen's or PAA's websites. Expensive media without sera or even w/o proteins are usually not necessary for a research lab. Cells can be adherent but only loosely attach. You should be able to suspend them by shaking the bottle. Usually cells should be split every 2-3 days. Medium colur is a good indicator for splitting time, too. Check the supernatant by dot blot for presence of the antibody and/or a functional assay with the corresponding antigen.
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PCR and Southern blotting differ per methodology and the majority of us here know these techniques in detail. But I was wondering why a researcher would prefer to conduct a Southern Blot over PCR, as both methods seek to identify a sequence(s) of DNA/RNA and as PCR has provides more advantages to offer?
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Some (but certainly not all) applications of southern blot that you cant easy do with PCR,
1. Detecting multiple homologous genes in a genome - PCR tends to be very specific and when two bands are seen, it is more likely non-specific or contamination
2. Detecting orthologous or paralogous genes in similar or distant species where you might not know anything about the sequence divergence and hence primer sites (you can increase or decrease specificity depending on hybridisation conditions) - if you dont know the primer sites, you cant do this
3. Detecting insertions of a plasmid/viral vector when making stable transgenics and determining copy number (similar to #1 really). Cant with normal PCR, qPCR you could
4. Easier to multiplex / detect multiple products
5. Similar to Kyle - you can analyse methylation patterns that affect restriction sites (as you digest with a restriction enzyme before probing)
6. Agree with Christian - replica plating - much easier/more time efficient than PCR
There is a couple of the top of my head. I am sure there are lots more out there.
How can I get rid of pigment contamination in my Trizol-isolated RNA, if neither additional column clean-up nor chloroform extraction helps?
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I extracted RNA from leech skin via Trizol and ended up having too much pigment (probably melanin?) contamination (260/230 < 1). I tried a subsequent clean-up over Qiagen columns, as well as multiple precipitations with Trizol/chloroform, but nothing helped.
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Do you precipitate the RNA in the process and your contamination is then in the precipitate?
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I use vectors containing whole genome of HBV for my transfections. To check viral replication, I get the supernatant of the cultured hepatocytes and concentrate the virus by centrifugation over 20% sucrose at 25,000 rpm for 4 hours.
I am wonderring if my percipitate has plasmid contamination due to cell lysis and plasmid release to the media. If so, could plasmids pass throught the sucrose along with the viruses?
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With "empty" HBV capsids from E coli, they pick up nucleic acid, even without the internal binding domain. So I'd guess that yes, you would co-purify plasmids and other junk inside the capsid if not also around it.
Dialysis of the resulting HBV band *might* remove it. (although you likely have enveloped and I'm not sure then...)
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What is the most RELIABLE method? From sample preparation to visualization, any suggestion for the N of such a study to get an acceptable error bar?
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I have had it quite rough with pCREB. I think its too fragile and really needs you to be quick when extracting the protein and mantain low temperatures as much as possible at every step. And make sure you have phosphatase inhibitor in your lysis buffer. The antibody (p-Ser 133) binding is very specific, so needs longer time (overnight, 4C!)
Also, I have just noticed it is better to add the loading buffer to the sample soon after extraction if you are going to store it for later use. I used to extract and freeze to -80 for later use and then add loading buffer just before loading for SDS-PAGE, but I seldomly got the desired bands!
I will be glad to see more tips here too! Am still working with pCREB and CREB western blotting.
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How can I get rid off some unspecific haemolytic substance in seed extract that has blood group specific lectin? The haemolytic property is neutralysed by adding bovine serum albumin, though but it is undesirable contamination to make it as reagent.
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There are some good suggestions provided to a similar question posed by Thierry Rolling on Dec. 20.
Also, if the hemolytic property binds to serum albumin you can try depleting it from your extract by passing it over immobilized albumin.
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Gel electrophoresis
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thank you for your question ,
please read this article:
Multiplex PCR for Detection and Typing of Porcine Circoviruses
J Clin Microbiol. 1999 December; 37(12): 3917–3924.
PMCID: PMC85845
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I have tried a number of extraction methods with no success, but I am not sure whether my technique is not good or there is insufficient DNA in the sample.
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Immerse the formalin-fixed samples in TE buffer for at least 2 hours or longer (e.g. overnight) before starting the DNA extraction? What is the purpose of it? What TE buffer will do?
what's the difference between siRNA and microRNA (miRNA) ?
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both are formed from dsRNA and both eventually get cleaved into pieces by Dicer and then incorporated into RISC which in effect cleaves target mRNA.
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Shortly: siRNAs (short interfering RNAs) are products of double-stranded RNAs, which can have viral origin or be products of repeats transcription etc. They bind to mRNA in the case of complete complementarity. miRNAs (micro RNAs) are products of dsRNAs encoded in genes of our genome. They do not require full complementarity to bind with target mRNA, e.g. one type of miRNA may regulate many genes, as well as one gene can be regulated by several miRNAs. For review, see, for example, Novina CD, Sharp PA The RNAi revolution. Nature. 2004 Jul 8;430(6996):161-4. or http://en.wikipedia.org/wiki/RNA_interference
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I have leaf-hoppers that I want to feed on an artificial diet (5% sucrose in TE buffer). How could I best design the feeding chambers using locally available materials, giving them enough room to hop and feed as well. I previously used 1.5ml microfuge tubes and the vectors died due to a lack of space. Anyone ever done this or anything related?
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I had an experience to feed the green rice leafhopper by 2.5% sucrose in a plastic vial (3 cm in diameter and 3 cm in height) covered with an artificial membrane of stretched parafilm. You can get more details from our paper “Insecticide Resistance of the Green Rice Leafhopper, Nephotettix cincticeps, to the Systemic Insecticides Used for Seedling-Box Application”
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I am aware of isolating exosomes from cell lines but I did not find much about isolating exosomes from explants. It would be highly appreciated if you share your experience on exosome isolation.
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We have had a great deal of success isolated exos from explanted tissue (breast, prostate, colon). Look at my paper from J urol 2010 for specific methods and let me know if you have any other questions
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Unfortunatly I already converted all RNA to cDNA... Could I still do whole trancriptome amplification? How can I overcome this problem of low cDNA volume and concentration?
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What do mean by whole transcriptome analsyis? A micro array or something else
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I heard nowadays isotype controls could be replaced by unstained samples.
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@ Ratnadeep: Please let me bring some answers for you:
1: Isotypes are not optimized for anything. They are non-specific Ab bearing the same fluorochrome as your specific one. Intracellular staining requires them EVEN MORE, as physical retaining of your specific Ab inside the permeabilized cell would go unnoticed without them. You HAVE to subtract this background, if not, you might see an increase in the signal that has nothing specific (imagine an Ab entering the permeabilized cell, but unable to be washed out after that: it will happen to the same rate in Isotype or in the specific).
2: You are right, but this information is always available somewhere from the company. Good companies put it in the technical data sheet, others put it in the lot specifications. If not, you have to call. But yes, it has to be equal. One is not to compare different concentrations of anything...
3: Theoretically, this could be appropriate. But in practice, it is risky. Let's imagine a simple case: stimulation of dendritic cells with LPS and evaluation of co-stimulation markers (CD40, Cd80, CD86). And, say... 3 time points. Nil, 4h, 8h, 12h. So you get your background signal for the makers in the Nil and you see an increase at 4h, 8h and a plateau at 12h (let's imagine). Then you think that all of that is specific, right? Deduce from the Nil and there you go. In a perfect situation: yes. But, what about the idea that the Ab receptors of the cell are also increased at the same time? You got blocking reagent for that, yes, but might be borderline... You might get many reasons for non-specific staining and not having you isotype control prevents you from properly quantifying. Once you have done it with isoptypes and controlled that the stimulation or time does not affect your staining, repeating without this isotype control is a possibility. But every new experiment has to be performed with its control (and should always anyway, but monetary considerations exist...).
Controls can be costly and annoying, but this is how to do valid and reproducible experiments. Having to repeat an experiment because other scientists will not accept a shortcut that you took is far more expensive than to do it right the first time.
Good luck!
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I want to express a few proteins in yeast and mammalian cells which should be translocated immediately to the mitochondria upon expression. Any vector or signal sequence information would be helpful.
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Agreed- if you add a signal sequence you can get it to go directly to the mito. For a matrix-targeted protein see my paper attached
Del Gaizo V, Payne RM. Mol Ther. 2003 Jun;7(6):720-30.PMID: 12788645.
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We biotinylate live cells (adherent monolayer of Ishikawa) for 1 hr at 37C. This is followed by quenching and washing with PBS. However after scrapping in PBS, followed by centrifugation at 2000 -3000 rpm for 15-20 min, cells do not settle down. It takes almost 30 min at 5000rpm to see a thin pellet. This is not the case with nonbiotinylated cells treated in the same fashion. Does biotin labeling alter buoyancy of cells? Is it ok to spin at 10,000 rpm or will it lead to removal of biotin tag from proteins?
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We use the biotinylated proteins for western blot. We opt to do whole cell lysate just after washing, to avoid this situation. The lysates will be cleared by centrifuge and equal amount of total protein was pull down with streptavidin beads, eluted with 2x sample loading buffer, and then run western blot. It is basically an IP procedure after washings. It works pretty well for us.
Why rifampicin does not bind RNA polymerase in eukaryotic cells?
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Action of rifampicin
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RNA polymerases (RNAP) from bacteria and eukaryotes are structurally very different. Rifampicin binds to the beta subunit of the bacterial RNAP which is very conserved among bacteria. However, the possible homolog of that subunit in the RNAPs of eukaryotes is not very conserved, which means they differ in aminoacid sequence enough to prevent rifampicin binding and inhibition. Summing up, rifampicin can't bind RNAPs from eukaryotic cells because they've mutated and changed a lot in those billions of years of evolution that separate those two groups.
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We just pulled a few interesting scFv (single-chain Fv) antibodies out of a phage display library. Now we want to produce a soluble antibody from these clones. While the phagemid allows for direct expression of tagged, soluble scFv in E. coli, we are also interested in reconstituting the scFv fragments in a way that ordinary, species-specific anti-Fc secondary antibodies will recognize them.
I'm aware that expression of full IgG is possible only in a eukaryotic system. I found a few protocols to make "humanized" IgG starting from Fab or scFv, but we want mouse antibodies. We are looking for a convenient vectors to fuse scFv to the murine Fc domain. Anyone know of a source?
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Dear Alexandra,
you might check out these two publications (PMID:17373908; PMID:17395713). They simply fused the scFv to different Ig Fc parts, lacking the CH1 region. Although they describe human IgG and IgE in this papers, they should also have vectors for murine IgG.