- Patrick Hörnschemeyer added an answer:How to concentrate proteins using Sephadex G-25 and dialysis?The situation is as follows:
Membrane protein elutes in acceptable purity after IMAC.
pH is 7.5, 300 mM NaCl, 10% Glycerin, 0.1% DDM
The problem is: Imidazole needs to be removed and buffer needs to be exchanged (for lower salt). And my protein needs to be concentrated.
Using Amicon Ultra or Vivaspin columns did not prove to be the right way. Concentration (100 MWCO) takes several hours and protein tends to precipitate after a while.
Buffer exchange using Superdex or PD10 columns works, but requires even more concentration afterwards.
I thought of this combination:
(dry inert) Sephadex G25
or very high molecular weight PEG (or something similar)
and Dialysis bags or devices with a MWCO of 100.000 Da.
So the idea is that I exchange the buffer while simultaneously concentrating my protein. But I guess it is only possible to dialyse first, and to concentrate afterwards.
The problem is that I need to make sure that I do not concentrate my detergent (detergent micelles of DDM are about 70 kDa in size and with my protein incorporated they are well above 100 kDa). Thus, requiring a dialysis membrane with a large MWCO, although people keep telling me that DDM is not dialysable. At the same time, this large MWCO would possibly allow the permeation of the "osmolyte" (PEG) into my sample. I guess this would not be the case with Sephadex, since the Particle Size is 85-260 µm.
Never tried the Sephadex G25, I would give it a try. With Spectra Absorbent it works super nice.Following
- Steingrimur Stefansson added an answer:Why do we mostly use only 6xHis Tag to purify Proteins Not 8xHis Or 10xHis?
When ever people make a clone with His Tag to their protein for purification by affinity chromatography why they mostly use only 6 His to protein. What will happened if we put more His suppose 8 or 10. I think by increase no. of His we can increase affinity of our His tagged protein hence chances of getting pure protein will be increased because it will elute with higher concentration of Imidazole.
@Alejandro, sorry, my mistake.Following
- Daniel Moran added an answer:Why does my protein concentration decrease after dialysis?I am purifying a 35 kDa protein, its temperamental and oddly, it crashes in low imidazole rather than in a higher concentration of imidazole. Anyways, I wanted to dialyse it for SAXS analysis at an initial concentration of 5mg/ml. However, after dialysis, when I took it to SAXS, and measured the concentration, it was 0.5 mg/ml. I can't really compute as in to why this happened. Anyone with ideas? Is this normal?
You could use a SEC chromatography resin to perform the buffer exchange but then the protein would need concentrated again. Acetic acid preciptiation or TCA of the same might work to concentrate or use a lyophilizer.Following
- Amrathlal Rabbind Singh added an answer:Can anyone help with protein purification?I have purified a protein of 21 kDa by Ni-NTA. The purified protein shows a single band on SDS-PAGE but three bands on Native-PAGE. The protein is a monomer, as confirmed by FPLC. The protein also seems to be aggregating as it does not even cross the half-way mark after the dye has run out on native gel. I need the protein for NMR and immunological in vivo studies and am worried of the two extra bands in further experiments.
My protein is not degraded by trypsin hence cannot do MALDI.
For MALDI you can use proteases other than trypsin (Arg-C and Asp-N) also for identifying the bands seen in the native gel.Following
- Adrian Abrahams added an answer:I am unable to decipher the gel side of the IPG strips purchased from Bio Rad. Can Anybody help in this regard?Earlier I had used Serva IPG Strips in which it was easy to decipher the gel side of the strips but recently we shifted to Bio Rad and here we are encountering this problem. Your valuable suggestions are required.
Did you manage to figure out the gel side of those IPG strips from bio-rad?Following
- Shujuan Gao added an answer:What are the recommended ways to preserve proteins that are to be subjected to downstream activity tests?
Hi everyone, may i know how proteins can be preserved upon purification? I need these proteins for downstream activity tests but as I will not need them right away, I need some solutions to preserve them. Is glycerol 10% with the elution buffer ok? By the way, I did His-tag purification. Thanks in advance!
There is no universal buffer can be use to store purified protein. First of all, you need to know some base information about the protein you are working on, such as does it require metal ion or other co-factor to stabilize, any conditions that might cause denature of the protein, will freeze/thaw cycle effect the activity etc, a lot to list, just mention a few for example. Some protein will denature when the concentration is low, you might use bovine serum albumin as a carrier, or precipitate with ammonium sulfate and just stored as is. 50% glycerol is a common choice. As long as the storage buffer you use does not interfere later on activity assay will be fine. Good luck.Following
- Akhil Raj P asked a question:Does anyone have experience in using profinity exact purification (Bio-Rad)?
For lysis of transformed cells which reagent are you using instead of NaCl??Following
- Dina Morshedi added an answer:Does anyone purify recombinant alpha synuclein?
When I look at the SDS-page pattern of alpha synuclein in some studies its band appears higher (around 17 KD) than the real place of its molecular weight (14.4 KD). However, our purified protein appears at around 15 KD. The sequence is OK and N-terminal also was analyzed and it is OK too. Dr.Sidhu in "Abnormal migration of human wild-type -synuclein upon gel electrophoresis" showed that ASN migrated in SDS-PAGE unusually, so why would our protein migrate and appear in its real position?
Thank you Sleman. In all reports it was expressed in E.coli which has not high PTM activityFollowing
- Andreia Ribeiro Albuquerque added an answer:Can anyone help with Toxoplasma gondii tachyzoites purification?Do you have any experience with the purification of TG tachyzoites from a culture of HELA cells? I'm planning to start with cesium chloride gradient or Percoll solution. Any tips or help?
Hello Roman, you have a couple good book chapters on-line (Roos et al, 1994; Striepen & Soldati, 2014) that speak of Toxo purification, as well as other nice infos about Toxo. Since their size is about 6 per 2 um, you can use a 3 um pore-size-polycarbonate-filter/47mm diameter (Nuclepore).Aggregated parasites and most host-cell debris remain behind. After that you can concentrate your sample by centrifugation and ressuspension of the pellet in whatever medium you think is apropriate for your experiment.Following
- Steingrimur Stefansson added an answer:How do we remove tween from protein solution?
Hi, we need to remove tween from a protein solution that will be analyzed by MS. What would be the available options starting by those cheap, effective and simple? Thanks!
In addition to the answers above, you can try these Pierce columns:
- Hemanth Kumar added an answer:Please suggest me why I am getting very less amount of my protein of interest compare to tagged GST protein after cleavage with Prescission Protease?
I am purifying a 8 KDa protein tagged with GST and cloned in pGEX 6P 2 vector. When I cleaved 8 KDa protein from GST by prescission protease I got very less amount of 8 KDa Protein compare to tagged GST protein. Cleavage reaction conditions are as follows- cleavage Buffer (50 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1 mM DTT, pH 7.0), Incubation time 4 Hours.
I am also attached gel photo for the same.
Please tell me why it happens?
How I will get more amount of 8 KDa protein?
try this.. reducing the incubation time and concentration of salt.Following
- Carol Ladner-Keay added an answer:Can anyone assist me with native gel troubleshooting?
I have question regarding Native gel (15%). My protein molecular weight is 13 kda and it supposed to form dimer when ligand is added.The ligand is around 500Da. When I run native gel, my protein dint enter into resolving gel.I ran it at 70 V for 7 hours in cold room and observed that my dye front ran till end,
Attached is my native gel picture. With reference to my native gel, the first 3 lanes are my protein alone. It is strucked up and not enter the resolving gel. Moreover I understand if the protein sample struck on the well, It means its getting aggregated. so the part of my protein is aggregated but rest dint enter the resolving gel.
The lane 4 -6 are the samples with protein and ligand. It supposed to form dimer when it run. But I couldnot interpret anything from the gel.
May I know how can I interpret results from native gel ? How can I check the dimerisation using native gel ? Which recipe I have to follow ?
I'm not sure, either protein degradation or a conformational change would explain a smaller band. Check your protein+ligand on SDS-PAGE. Does the ligand have a positive charge? I'm not sure what you mean by "just after it enters the stacking gel". When I run native acidic gels I run them for about 75 minutes at constant current of 30 mAmp. Hope that helps.Following
- Levani Zandarashvili added an answer:How to determine peptide concentration that is devoid of Tryptophan, Arginine, Lysine, Phenylalanine cysteine and Tyrosine?I have a peptide sequence with a molecular Weight approximately 4.5 KD. They don't have any Tryptophan, Arginine, Lysine, cysteine, Phenylalanine or Tyrosine. Can anyone suggest me a method to determine this type of Peptides concentration?
you can use amino acid analysis which is described here:
at some universities they do this in analytical chemistry labs.
This method worked perfectly for my 30aa peptide which had only acidic amino acids.Following
- Jose L Chavez added an answer:What causes the resin of Ni-Agarose column to attach to itself and make globular objects?
Is there any solution for removing these globular objects?
If yiu are using the ni beads more than once. This is a very bad practice and most of the times the beads are in that bad conditions that forms clumsFollowing
- Yaghoub Safdari added an answer:Does anyone have an idea as to why I am unable to get the correct size of human protein cloned in a pET28a vector?I have cloned a human gene into the pET28a His tag vector but I am unable get the correct size of my protein. I have confirmed recombinance by sequencing and restriction digestion.The correct size of my protein is 55 KDa but I always get a band of 75 KDa. Please give me suggestion on how to get the correct band?
I believe that neither His-tag nor post- translation modifications (Glycosylation and ect.) are responsible for the difference (20KD) you observed.
From the first ATG codon, which exists on the plasmid map before the first His-tag, to the stop codon, which lies after the second His –tag, will potentially lead to a molecular weight of only 6.8 KD ( approximately 1/3 of the difference you observed).
Despite their several advantages in producing recombinant proteins, bacteria are unable to accomplish the required post-translational modifications.Following
- Abhas KUMAR Maharana added an answer:Does anyone have an idea of a carbon source for cellulase excluding: orange and pineapple?Purification of cellulase from a sourceFollowing
- Zaiddodine Pashandi added an answer:Can you recommend a protocol for recharging Ni-NTA aGAROSE resin?
I use Ni-NTA Agarose resin to purification a protein with his-tag tail. As I use it multiple times, it seem that it has lost the efficiency. I want to recharge it to become refreshed.
What is the best protocol for this?
Thank you dear EllisFollowing
- Paul Lesbats added an answer:Can you recommend a good GST inclusion body solubilization and renaturation kit?
I used GST gene fusion system (pGEX-6P-1) for protein expression and found all of my recombinant proteins deposited in the form of inclusion bodies. I changed the conditions (IPTG conc, induction time, tem) but no advantages, all remain in inclusion bodies. So I would like to solubilize and refold my proteins. Is there any good commercial kit available for this purpose?Following
- Gary Laco added an answer:How could I elute my protein from Ni-NTA beads when do purification?
My protein with His tag is always on the Ni-NTA beads when purification and can not be eluted.So, how could I solve my problem? Thank you for your advice~
Use 10-fold less beads and 10-fold more protein. You need to first saturate the hydrophobic binding sites on the media, and then the Ni sites, then you may be able to elute your protein.Following
- Rao Kv added an answer:Can someone advise on availability of codon optimised cDNA Clones for Malaria MSP-1 antigens 19Kda protein?We are in need of cDNA clones for the Malaria MSP-1 Antigen of Malaria P. Falciparum and P.Vivax for our development works. Vector ligated or cDNA clones of the above MSP-1 protein are of our interest.
Dear Mr Anil, Thanks for your reply as such the cDNA clone is meant for diagnostic work and we need to produce from E.Coli. Off late the resolution to my interest is also found from the companies in India and abroad. Thanks for your forwardness on my quiries. Have a great week ahead and nice to see you on R.G.Following
- Sanjay Mishra added an answer:Can someone please explain why there are 2-3 bands of my protein after purification?
I realized that this often happens only when I perform purification. My protein is fused to a 60kDa tag and my protein it self is ~20kDa. So taken together, the fusion protein should result in ~80kDa band. When I express this protein, it is a single band. However, when I purify the same protein on the Nickel packed column, more than 1 band (bands below the red arrow) is eluted out. What could be a plausible reason for this?
I hate attached the purification image here.
In my opinion, it is very common in this context. What exactly, you are advised to work on with probing with specific antibody followed by 'Western Blotting' technique. you may probably obtain the good results likely to be reproducible. further queries are most welcome.Following
- John T Corthell added an answer:What is the best way to purify phosphorylated proteins from brain tissue?
I want to look at phosphorylated proteins purified from mouse hippocampal area CA1. My initial though was to micro dissect the CA1 from the brain and then flash freeze it in liquid nitrogen, before continuing with cell lysis etc. I'm worried the dissection might take to long and since I will be looking at phosphorylations, time is at the essence.
Hope you can help with some suggestions.
Do you have any phosphatase inhibitors? You can make a solution of sodium orthovanadate, for example, and include that in your protease inhibitor mixture. Including inhibitors in your mix will make this a lot easier.
I can usually get tissue out fast enough to avoid a huge loss of phosphorylation. Flash-freezing, like you suggested, followed by keeping the tissue at -80 degrees C until homogenization, usually works just fine. Don't leave the samples in the freezer too long; I've had some degrade in the -80 after 6 months or so.Following
- Xiaoqin he added an answer:Lysis of SF9 cells infected with baculovirus to extract target protein for further purification?Recently I started to work with insect cells to produce protein samples for structural determination. I have experience of protein purification from E.Coli but I have a few doubts about the extraction of the target protein from insect cells. In a typical purification, starting from a pellet of insect cells obtained from 2.5 L of culture, I suspend the cells in about 70 mL of lysis buffer (50 mM TRIS pH7.7, 300 mM NaCl, 5% glycerol, 0.2 mM PMSF, protease inhibitor tablets) the lysis is performed adding Triton-X100 to a final concentration of 0.1% and incubating for 15 minutes on ice, mixing gently every 3 minutes. After that a mild sonication is performed (2 seconds “on” 10% amplitude/60 seconds “off” repeated 15 times). The mix is then spun down at 20000 rpm for 30 minutes at 4C to remove insoluble particles. At this stage my lysate looks very cloudy and it is difficult to filter it with 0.45 um syringe filters. Moreover, a layer of white “stuff” is also visible on the surface of the lysate. Eventually, after changing lots of filters, I managed to filter the supernatant and the protein is successfully purified. I believe that I lose about 30% of the total sample in the filtering process of the clarified lysate. Am I doing anything wrong? Could anyone suggest how to fix this problem?
Hi, if you think filtration is the procedure that you are concerned, why you don't centrifuge at higher speed before filter the lysate? I usually do ultra-centrifugation at 120 000 g for 20 minute, the insoluble material that you observed should be spun down.
- Alejandro Gonzalez added an answer:Hoe can I best carry out binding, washing and make my own elution buffer?
I was using column base protocol, but I don't have more buffer for it, but a lot of columns so, how can I make my own buffers?
This is because I need to purify my DNA and I don't have time to wait the company to send me another Purifying Kit so, I want to know if its easier to do my own buffers in my Lab.
Thank you and have a nice day! :D
Wooaah!! excellent! thank you both for the fast and helpful answers!
- Sai Shyam Narayanan added an answer:What are the safe conditions for sonication procedure in E. coli cells?As you probably have noticed, sonication of E. coli has several risks for our recombinant proteins. This is due to the ultra-sonication procedure, through cavitation, can rise the temperature quickly inside the cell suspension. Therefore short pulses are preferred over a long continuous pulse and it is very important to maintain the cell suspension in chilled ice during sonication. Otherwise, sometimes aggregation can be induced by this procedure in your target protein triggering to insolubility.
Maybe there is not a perfect sonication procedure which can be used for all the recombinant proteins, possibly it will be depend on the expression levels of the rec-protein, but I think that we can find out a condition range in which all the people working with recombinant protein performed in E. coli could work without too much risks.
I would appreciate very much if you share your knowledge in this controversial issue, sharing your sonication conditions. Sometimes in biblio, conditions are not well defined, it is necessary to know: type of machine, diameter probe, cell suspension volume, vibration amplitude and % used, time pulse on/off and the whole time procedure. In my case:
SonoPlus, 3 mm diameter probe (with 210 um amplitude capacity), 20% vibration amplitude, 5´´ on/5´´ off for 30´´ (x3) with 10´´ rest between 30´´ cycles, 5-10 ml cell suspension.
Any other hints will be welcome!
I think it is important to standardize sonication condition for consistency in the protein yeild, purity and monodispersity. I have no clear evidence for this statement, but my experience say so. As yo rightly pointed out there are no good refernces providing the possible problems one could face for not having followed an optimal sonication conditions. Scaling up issue in sonicator has also been challenging. I use 6 cycles of 20 sec ON/ OFF with 50-60% amplitude and 50% duty cycle on ice for 3 mm probe. But scaling these conditions to a 9 mm probe did not give similar cell lysis. Could you discuss about how sonication steps are scaled up with consistency ??. Normally I have seen my colleagues using French press for higher cell mass. Can sonication with bigger probe (diameter) replace this ??.. Does the output voltage displayed in the machine always speaks for the consitency ?Following
- Anthony Harrington added an answer:Can anyone help with purifying crude protein extracts?
Hi, I have a question about crude protein lysates (extracts). A fellow lab member in my lab is trying to isolate a 27KDa protein from here crude protein lysate. She mentioned that this is the smallest protein in her lysate (confirmed with SDS-PAGE). Right now, she is planning on running gel-filtration to isolate it but has yet to start. She did ask other labs at the school that have HPLC/UPLC about running her samples but it seems that something always comes up right before its time to run the sample. I mention that she could use a membrane 30KDa MWCO filter along with a vacuum filtration set-up to isolate her protein. I was curious if this is possible? She is ignoring my suggestions and I just want to know if its because my suggestion makes no sense. Any advice will greatly appreciated and will be passed on to my lab mate.
Update on results, she has ran the size exclusion chromatography and got inconclusive results. The void volume fraction has the same bands as the sample not passed thru the column. She is using biogel P30, with tris buffer at pH 7.6. The fractions after the void volume don't show any bands. This is her first try at this method. She is looking for any suggestions that might help her troubleshoot this issue. I am busy with my own project so I cannot provide any assistance to her except by passing along this information and fowarding any suggestions provided. ThanksFollowing
- Dongfen Yuan added an answer:Can anyone help with hydrophobic interaction chromatograph for protein purification?
My protein has better absorption at 220 nm than at 280 nm. So I am going to use 220 nm since my sample may be really diluted. I am using gradient elution. Mobile phase A: 1 M ammonium sulfate + 50 mM PB, pH 7, mobile phase B: 50 mM PB + 40% isopropanol, pH 7. The absorption at 220 nm goes down as mobile phase B increases during gradient elution, while absorption at 280 nm is flat when nothing or only buffer was injected. The shape of absorption at 220 nm looks like the shadow of the gradient curve. Does anyone has any suggestion?
Thanks Adam. That is my plan now.Following
- Russell Jarrott added an answer:Why nikel ion does not bind to Ni-NTA column after APS treatment?
I treated Ni-NTA column with 10% APS because I purified a protein using a buffer containing DTT and it tourned brown. After APS treatment I wash the column with water and then I stripped it with EDTA, wash it again and finally used NiCl2 to recharge column, but when I washed again with water all the nickel was eluted. Is the column damaged?
I would assume your column is damaged. I agree with Dawid that 1% sounds more suitable. The snippet I found about APS cleaning did not actually state the %. If it hasnt bound the Ni again then something has obviously changed. However there are also different levels of Ni on resins. ie GE/Roche are a bright blue, PrepEase is almost white. So there may just not be as much bound as previous. But if it doesnt go back to 'normal' I wouldnt trust it.
I was more addressing the need that caused you to ask the question. If I want to use high DTT (5-10mM) then I wash with 500mM Imidazole first (or after) and this seems to fix the problem of the resin turning brown.
- Gergely N. Nagy added an answer:How can I get rid of air in an AKTA purifier inlet tube?
Accidentally air was sucked in by the pumpwash protocol as the pump inlet was not immersed in solvent. Currently the pump is not capable of sucking up solvent whaterver hard it tries. Could you please give me an advice how to re-introduce solvent in the tube system? Thanks in advance!
Hi Misha, Matteo and Heide!
Thanks for all the comments! I these tricks on the instrument on monday when i will have access- and report here how it worked.
I am really indepted for your advices!