- Adam B Shapiro added an answer:2Any advice on purification peptide coupled-quantum dots?
We are doing EDC coupling of peptides (around 8 kDa) to quantum dots (around 6 nm). We would like to separate the conjugated QD from the unbound one. In your experience, what is the most recommendable method? We tried spin ultrafiltration with a 10 kDa column also to eliminate the free peptide, but the QD remains bound to the membrane, and therefore we are not able to separate the conjugate from unbound QD. We were thinking about Sephadex or SDS-PAGE as well. But may be the recovery is significantly reduced if we perform SDS-PAGE...
Thanks for any advice!
There are potentially 3 categories of products: (1) peptide-coupled quantum dots, (2) uncoupled quantum dots, and (3) uncoupled peptide. It should be easy to separate (1) and (2) away from (3) because of the large difference in size, using gel filtration, centrifugation, sucrose gradient ultracentrifugation, dialysis or ultrafiltration (if the non-specific sticking problem can be solved).
It would be more difficult to separate (1) and (2) because they are so similar in properties. The need to do this could be avoided by making sure all the quantum dots have peptide attached by using a large molar excess of peptide over quantum dots. However, a possible approach to the separation would be to make the peptide with an attached affinity tag, such as hexa-histidine or biotin. Then, an affinity resin such as immobilized Ni2+ or streptavidin, respectively, would be used to pull down (1), leaving (2) behind, Finally, (1) would be eluted from the resin by imidazole or heating+biotin, respectively.Following
- Tang Xiaoqiang asked a question:NewWhat could cause a failure to detect proteins with SDS-PAGE and Western blotting from baculovirus-infect insect cells?
I transfected low-passage healthy Sf9 cells with PCR-verified bacmid(Bac-to-Bac system) and saw all the signs of infections during transfection and baculovirus amplication of P1 and P2,but I could not see the specific band with Western blotting and SDS-PAGE for P2 and P3 and P3 virus-infected(with varying MOI and time) cell lysates(cells were harvested when 70% was dead,as suggested by the manual).Thanks.The Bac-to-Bac manual says"Viral stock contains a mixture of recombinant and non-recombinant baculovirus",however,the PCR of purified Bacmid gives only a specific band of expected size,without a 300bp band.Can anyone give me some suggestion?Following
- Panduranga Rao added an answer:2Why E.coli produces both fusion protein and fusion protein+recombinant protein?
When I produce recombinant protein by expression vector such as pMAL or pET43
Look for possible mixed culture (recombinant and non-recombinant). Pick a SINGLE colony and test it again.Following
- Ade Kujore added an answer:3How do I regulate the flow in an ion exchange column?
I am trying to isolate a nerve growth factor, and the protocol calls for various flow rates throughout the process. I have a really old flow adapter in the column, but I really cant seem to regulate the flow to match the protocol.
Is there anything I can do? Perhaps a pump that regulates the flow?
Are you using a glass column, how is liquid passed through the column. What is your detection technique. Please provide more details.
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- Ahmad Fazeli added an answer:4How can I purify a recombinant protein without urea and salt?
I had produced and purified the recombinant protein by genetic engineering. However, the protein was eluted from resin with high salt and urea solution. When I use dialysis method to remove them, the proteins aggregated. When using ultrafiltration, the protein did not accumulate in upper solution. I just want the protein in low salt solutin but with high concentration.
There are several nonioni surfactant e.g. Polysorbates, and, plutonics which you can use during your buffer exchange dialysis/UF process.Following
- Jorge Ruiz Carrascal added an answer:4Is there any quantitative assay for protein degradation study ?
During the study of protein degradation, Substrate and product quantification using conventional methods like Bradford , Lowry etc. could not applied because they are based on the reaction of added chemicals with amino acid having Benzene ring in their structure. And also the bacteria which are responsible for protein degradation release enzymes for protein degradation. These enzymes are also proteins.
Although there is qualitative assays are in literature but quantification of protein degradation with respect to growth kinetics and enzyme kinetics are lacking. So any one can suggest me How can quantification of bacterial protein degradation possible?
A very common method for following protein degradation in some food products is quantifying total nitrogen, peptide nitrogen and amino acid nitrogen separately (or even just total nitrogen and non-protein nitrogen): it will give you an idea of how intact are the proteins and how intense is the degradation. The quantification of nitrogen is through Kjeldahl.
- Neha Arora added an answer:3How do I purify an insoluble protein present in a pellet?
I expressed protein and unfortunately it became insoluble and remained in cell pellet. How do I get protein from pellet, so later I can use it for antibodies production?
Hello Prem! There are a number of detergents widely used to solublize protein. I can tell u some detergents: trition, CHAPS, sarkosyl, urea. Generally, sarkosyl works well,however it is a harsh detergent and may affect your protein activity. In case you use it try to optimize and use as less as possible to get enough protein for purification.Also make sure you remove the detergent by dialysis, so that it does not hamper your further applications.Following
- Alejandro Martin added an answer:6What is the protocol using ammonium sulfate for purification?
Had anyone here tried out using ammonium sulfate to purify your protein? How are you actually work on it? Carry out using affinity purification first then salting out the remaining impurities? Anyone mind to share your experience on how you actually optimize the concentration and use it for purification? Anyone has protocol for reference? Thanks!
Yes, I've done it before. First you add ammonium sulfate at a specific saturation % to precipitate proteins other than your target protein, then add additional ammonium sulfate at a higher saturation % to precipitate your protein while -hopefully- leaving other contaminants behind (provided that the protein does not crash irreversibly in this step). The exact % have to be determined experimentally beforehand; go google a bit and you'll find better descriptions than anything I can provide.
The point is -and judging from your comments, you are well aware of this- that AS precipitation is not very selective. You use it in a crude lysate and it gets rid of a lot of contaminants, but will never yield a 'pure' protein by any sensible criterion, and the reason is that the minimum AS saturation % at which all of a given species will precipitate is not a single well-defined value, but a range that often overlaps with that of other protein species. If you use it after an affinity step it may improve the purity of the resulting preparation, but will never work as well as another run on -say- a gel filtration column.Following
- Alejandro Martin added an answer:7What can I do to prevent my protein from precipitating on a Nickle IMAC column during denaturing conditions?
I'm trying to purify an over-expressed membrane protein which I suspect is stuck in inclusion bodies. I extract the proteins using 8M Urea in a 50mM Tris-HCl (pH 7) buffer containing 20mM imidazole and 0.5M NaCl. I ultracentrifuge the sample before loading it onto the IMAC column. During loading, the column gets clogged quite soon. When I try to elute the protein at very low flow rates on FPLC (to prevent over pressure), the proteins come off very slowly (during most of the linear gradient from 20mM to 500mM Imidazole), especially my protein of interest.
What should I try to prevent the protein from precipitating/clogging onto the column?
Can something like a phosphate buffer possibly have a different effect vs a Tris buffer? Should I try a higher/lower pH? Less NaCl?
PS: pI of my protein is 8.5, truncated version of the protein (just the hydrophilic part) purifies perfectly as it's completely soluble.
Try passing the sample through a 0.22 um filter. If it clogs the filter, it will definitely clog the column.Following
- Sreenadh S Pillai added an answer:7What is the best buffer for MMP-2 enzymatic action?
I am using MMP-2 to cleave the GPLG-VRGK short peptide. I have tried it in water? Does MMP-2 is active in water? Could you please tell me why it is not active in water? Which will be the best buffer for MMP-2 and why? What is the role of buffers in proteinases substrate specific activity or enzyme activity?
Please be kind enough to give me your advice.
Thank you very much Dr. Marcia Moss for the kind information. I have read your few articles will be helpful for my future research.Following
- Minta Wang added an answer:3What it is the best tag for purifying recombinant proteins in mammalian cells?
I am looking for a tag that allows to purify recombinant proteins in mammalian cell with high affinity and high specificity in non-denatured condition. Can someone advise me what it is the best tag available for my need?
Flag tag has a good binding specific. If large tag is acceptable, FC tag may be a good choice.Following
- David Farringdon Spencer added an answer:3How do I assess the DNA contamination of a protein preparation?
Can we trust the 260/280 ratio?
Do you have an alternative method to propose?
- Sebastian Schmitt added an answer:1Number of HEK293 required for pull down followed by gel filtration
I am studying a protein involved in DNA damage pathway. For my experiments, I wish to treat HEK293 cells with genotoxic agents: bleomycin and florouracil and UV.
My protein of interest is halotagged (for pulldown) after which I wish to use gel filtration to get to separate any protein complexes that may be formed and how they vary in each pull down.
However, I do not know how many cells I should start with to get a good separation on the gel filtration column. any ideas.Following
- Paul C Engel added an answer:6Is it better to refold a protein bound to a column or free it into solution? I'm refolding by lowing down urea concentration. The column is a NiNTA.
I'm trying to get the apo version of my protein. I get rid of the ligand washing with 3M of urea but now I need to refold it without misfolding it. Not sure if it's better to refold bound to th NiNTA column or free in solution by dialysis.
I would be less optimistic than the other contributors about this approach, because I think there is quite a big chance that, on or off the column, successful refolding may depend on the presence of the ligand you are trying to remove! Could you maybe try a gentler way of removing the ligand - i.e. without complete unfolding? You could use either lower concentrations of denaturant to loosen binding or dialysis with activated charcoal in the external buffer, very effective in removing nucleotide ligands for example.Following
- Paul C Engel added an answer:5Is it suitable to carry out centrifugation in normal centrifuges (instead of cooling centrifuge) while doing enzymatic assays?
Hello, I wish to know Whether it is good to carry out centrifugation in normal centrifuges instead of cooling centrifuges while doing enzymatic assays especially in cell culture.
In general, if you are dealing with extracts of human or other mammalian cells, it is likely that cooling will be important in order to slow down proteolytic degradation.. However, the best procedure is to assume nothing and actually test the requirements of your own system. If you can show that the protein you are working with is stable at room temperature over the relevant time period then you may be OK. There is also the possibility (and this is a very strong reason for CHECKING and not just assuming that you have to do what everyone else does) that you might have an enzyme that is COLD-LABILE - stable at 37C or maybe at room temperature but not at 4C. It happens! So try out the different conditions for yourself.Following
- Mauricio Hernandez added an answer:15Why I didn't get any bands present on SDS PAGE?
My experiment is about protein expression in E.coli.
Done with growth, cell lysis and purification.
Failed in my Sds experiment.
Only the protein ladder gave the band, my 2 sample and positive control failed to give the band.
To quantify proteins ?, chances are you've loaded very little.Following
- Wajd Amly added an answer:3How can I distinguish between Primer dimer vs non specific amplification?
How can we distinguish between Primer dimer and non specific amplification using gel electrophoresis and melt curve?
Thanks a lotFollowing
- Brandon Ambrose Kemp added an answer:4What is an efficient procedure for isolation of plasma membrane proteins from cultured animal cells ?
I would like to check wheather my protein is located in plasma membrane. Can anyone recommend an efficient procedure for isolation of plasma membrane proteins from animal cells cultured in vitro? The protein of interest will be identified using Western blotting method. Thank you for any help you can provide for this question.
you can perform a surface biotinylation procedure.Following
- David A Johnson asked a question:NewParamecia in situ PCR?
I am attempting in situ PCR with paramecium. Has anyone tried in situ PCR in a similar organism with a pellicle? I trying to determine the parameters for fixation and protease treatment. Any help welcomed.Following
- Lindsay Becker added an answer:4How do I stop RIPA-insoluble, Urea-soluble protein fraction from precipitating in Western blot wells?
I’m trying to analyze insoluble proteins from mouse brain. Basically, I extract with RIPA, then take the pellet that’s left over and extract with a urea buffer (7M urea, 2M thiourea, 4% CHAPS, 30 mM Tris, pH 8.5).
When I do a Western blot, the RIPA fractions run perfectly, but some of the wells with urea fractions streak out on the blot (lane 6, ponceau #1 attached, all lanes contain urea fractions from different mice). The streaking occurs from when the proteins first start to enter the gel. The most of the loading buffer is pulled into the gel, and some of the protein stays in the well in trickles out in a line from the center of the well.
One time, I loaded each urea fraction in 2 different gels at the same time. A fraction would run fine in one gel, and then the exact some fraction from the same tube would streak in the other gel in the same rig.
I’m using 4-12% Bis Tris NuPAGE 15 well gel, 1X MOPS SDS NuPAGE running buffer, and NuPAGE LDS sample buffer. I run at 100V for 30 min then 150V for 45 min.
After playing around with different ratios of protein in urea, distilled water, and 4x LDS sample buffer, it seems 5 uL of each of the aforementioned seems to work best. However, I’m still seeing some streaking, and the even lanes that aren’t streaking don’t seem to be running perfectly (ponceau #1).
Other things I’ve learned:
If I bring the LDS sample buffer to 2x final concentration, the loading buffer looks like it’s coming out of the sides of the wells as it enters the gel and there is strong banding on the sides of each lane (ponceau #2 attached).
Bolt gels are absolutely terrible. Every lane streaks out when I run the samples on a bolt gel.
Any advice or guesses as to what may be happening would be greatly appreciated! Thanks!
I'm not heat denaturing the proteins because they're in urea. Heating in urea causes carbamylation and degradation of proteins, and it degrades the urea.Following
- Atilio I Anzellotti added an answer:4Do you know a practical and industrial case study of protein mixture for separation of proteins?
I will want to separate and purify protein mixtures by liquid chromatography method in simulated moving bed (SMB) based on difference of surface charge of proteins. Do you know protein mixture as a industrial case study that have this property?
Gel Filtration Standard #1511901 from BioRad gives you a mixture of five proteins, it is better for size-exclusion HPLC though.Following
- Deepan Shah added an answer:15How best can I refold a recombinant protein from inclusion bodies and retain its bioactivity?
As a novice in this field, I would like to obtain a bioactive refolded protein from inclusion bodies. My his-tag protein is 77kDa cloned in pET32a (+), pI 5.84 and contain 8 cysteines. I tried soluble expression with low temperature (10-20⁰C), different IPTG concentrations, changing vector (tried pGEX-4T1), and modifying LB media with additives but all yielding to inclusion bodies. So i decided to solubilize the IB and try to refold. I successfully solubilized in 2M urea pH 9 with 2mM TCEP or 2M urea pH 7.5 with 1mM DTT. I have tried to refold by either gradual removal of denaturant (dialysis) or rapid removal (dilution) but still I cannot have bioactivity. Refolding buffer composed of GSH/GSSG(10:1), 1XPBS, NaCl , EDTA and L arginine. Later I tried replacing PBS with 50mM Tris-HCl. Can someone advise me on how to resolve this issue. Thank you in advance.
I meant TCEP in the refolding buffer.
It looks like you have a zero refolding yield at present. Try some of the things suggested by your correspondents here and see if that improves. If there are any cofactors for your protein try adding them to the refolding buffer - sometimes proteins that are heteromeric need their partners to refold correctly.
I don't know anything about antioxidant proteins in particular but if you check whether your protein or other structurally similar antioxidant proteins have been refolded previously by searching the literature, you may find the answers.Following
- Ulrike Jung asked a question:NewWhich large scale Tangential Flow purification system is recommended for vector purification?
Is anyone out here who has done vector purification with TFF? I'm looking to compare experiences with systems, especially anything above 1 liter. What did you use, what were your experiences, which system would you prefer?
And if you used the GE AKTA flux systems (S or 6) or maybe even the Spectrum Pilot Plus some input on how they perform would be fantastic!Following
- Subhas Chandra Bera added an answer:5How can I purify a protein using guanidium hydrochloride?
I am purifying a protein with a his-tag using 6M guanidium hydrochloride. At what step the concentration of guanidium hydrochloride should be gradually reduced, during bead binding or during washing steps?
I also agree with Kurt D. Bernd. Otherwise you can elute the prtoein in denaturation condition and then concentrate it with Amicon filtration in the same medium. After that the concentrated protien can be dialysed in gradinet denaturent buffer from 6M to 0M at required temperature. Good luckFollowing
- Rafik Karaman added an answer:1Can someone suggest a suitable FTSC fluorescent label of proteins?
Could someone suggest a protocol for protein label, i'm using a protocol on which i wash the protein with ethylacetate-ethanol after Precipitation of protein with 20% TCA, i have a doubt that i lost my protein after the washing step, could i skip this step ? or do any thing else, replace it? any suggession please? Thanks,
You can use the following:
Thermo Scientific Pierce Dye Removal Columns effectively bind to unconjugated fluorescent dye molecules from protein solutions to rapidly purify fluorescent conjugated antibodies and other proteins after labeling reactions.
These Fluorescent Dye Removal Columns enable fast and efficient removal of non-reacted fluorescent dyes from protein labeling reactions. Removing excess dye after a labeling reaction is often difficult and time-consuming but is essential for accurate determination of dye-to-protein ratios. The dye removal resin in this kit is highly specialized to produce exceptional protein recoveries while effectively removing non-conjugated dye. Using the appropriate amount of resin and buffer conditions, almost any fluorescent dye can be removed with this kit.
Features of Dye Removal Columns:
• Fast—removes free, unconjugated dye from labeled protein solutions in less than a minute
• Specific—purification resin provides outstanding conjugate recovery (75 to 95%)
• Customizable—separate resin and microcentrifuge columns allow optimization for different dyes and concentrations of sample
• Sample-friendly—processing results in minimal sample dilution
For more details please use the following link:
Hoping this will be helpful,
- Syed Zahid Ali Shah added an answer:6Protease inhibitor coctail quantity in brain homogenate?
If I want to purify prion protein scrapie (PrpSc) from brain tissue of mouse and in the first step I need to put protease inhibitor cocktail in PBS, then homogenize brain in it. I have 100x protease inhibitor cocktail but I am not sure how much I need to put in per ml of PBS?
I would appreciate someone's help please.
thanks to both of uFollowing
- Anton Beletskii added an answer:2Any advice on why KLF2 is not appearing on Western Blots?
We are trying to detect and quantify the presence of KLF2 which should appear as we have memory cells (CCR7+/CD62L+). We're using western blotting for that and we've tried both the traditional "long" method, and we've tried ProteinSimple's Wes Machine. Neither can produce a confirming band.
-We've tried multiple antibodies from different companies
-We've tried checking our cultured cells from D0 through D6 every day to see if KLF2 may have come and went
-Maybe we thought that KLF2 exists in small quantities, so to exclude the noise from extracting all the proteins, we've done a nuclear extraction and ran a western blot on that- No band
- We've tried varying both input protein concentration and antibody concentration
Any suggestions would be great! We've been trying to detect this elusive KLF2 for a while.
We've ran western blots on many proteins, and those work fine. Only KLF2 doesn't work for some reason...
I am not an expert in this particular target, but it seems that proteasome inhibitor Velcade is boosting levels of KLF2.
You can try to boost KLF2 levels by treating your cells with Velcade, or at least treat them with protease inhibitor cocktails (cOmplete) during protein isolation to ensure integrity of KLF2.Following
- David Lubkowicz added an answer:2Is there anybody who works on expressing exogenous protein in LAB?
Recently,I am expressing an exogenous protein with size of 450bp in L.Lacits NZ9000 using pMG36e. The protocol from High-Frequency Transformation, by Electroporation, of Lactococcus lactis subsp. cremoris Grown with Glycine in Osmotically Stabilized Media(PMID16348073 )isreferenced.Unfortunately,I can't get positive clones after electroporating ligation mixtures into competent cells of L.Lactis NZ9000. Can you help me with this problem? Thanks very much.
That method you are using is actually quite efficient. I used it to transform lactis successfully.
What is your transformation efficiency when you just transform pMG36e (just the original plasmid)?
Are you using the suggested SR plates?
If that is reasonably high, like 100+ colonies for 100 ng plasmid then you can try the ligation mix.
Here a couple of suggestions regarding troubleshooting:
- The ligation mix has to be purified. Either using a PCR purification kit or a phenol extraction as the Ligationbuffer etc reduces efficiency strongly
- Verify your ligation via PCR (1 primer insert, 1 backbone) to confirm that the ligation is step is successful
- Electroporate at least 1 ug of purified Ligation mix and plate everything
Hope that helps.
- Jaime Guillén added an answer:3What type of MWCO Membrane filters used to concentrate 10Kda protein?
Membrane filters to concentrate 10Kda protein
You must use filters MWCO<5000. There are a lot of commercial suppliers, Sigma, Millipore, Sartorius... and different membrane types. The most common used material is regenerated cellulose. But, sometimes for some proteins is better use other type of membranes in order to avoid precipitation; polyethersulfone (PES) or cellulose triacetate (CTA) for example.Following