- Eric Turk added an answer:3What factors can affect the dissociation of hemoglobin in to dimers?
I am using gel filtration chromatography to analyze Hemoglobin at varying pHs. The results of the GFC at a pH of 7.4 showed that hemoglobin was in its dimer form (I was expecting to see Tetramer). Is it possible that there are other interactions affecting the elution of the hemoglobin in the column?
My experience was same as that of Kristian Stødkilde-Jørgensen. An interesting phenomenon I found surprising, since texts refer to tetramer exclusively.Following
- Eric Turk added an answer:5Strange Gel-Permeation Chromatogram?
my latest gel permeation purification produced some strange results (see chromatogram attached).
I am purifying a small 14 kDa protein after Ni-NTA-affinity chromatography and subsequent tag cleavage (TEV). I am using a Sephadex 26/600 column (GE) at 1,5 ml/min. Sample volume is 15 ml.
I added Blue-Dextran (2 MDa) to the sample in order to have an orientation concerning size.Now the first elution peak is therefore definitely Blue-Dextran, but what about the other two unshapely signals? (Blue line = OD280nm)
My column is rather old, and the sample is not transported evenly, but is somehow distorted (like a 45 degrees tilted disk) - this might explain why there is such pronounced tailing? Since my protein is aggregation-prone, the middle peak might arise from the dimer (with severe tailing).
But what puzzles me the most is the strange increase in conductivity (red line) at the end of the separation. How can this happen? I am clueless, since the column was perfectly equilibrated in running buffer... Also the respective fractions having this strange effect do not have the pH of the running buffer (pH 4.5) but the pH of the sample (neutral) - what happened?
Plus: Is the separation sufficient? What could I do better?
Any help on this case is gretaly appreciated!
Several mutually-reinforcing clues in your image and description: The 45° tilt you describe (presumably referring to the blue dextran band visible through the glass cylinder) indicates either sample viscosity is too much greater than that of the running buffer, or there is precipitated/clogging protein or debris/bacteria from previous runs asymmetrically obstructing the topmost region of the column, or poor column packing (least probable since this column was sold professionally pre-packed). The Blue dextran peak should not have a shoulder; the same for the salt conductance peak at the end. Your peak of interest is behaving like these, but since in the better MW resolving region of the chromatogram, the trailing shoulder is more elongated. Note that sample viscosity is increased by protein concentration as well as small solutes. Protein precipitates in the column are effectively removed by running in 2 column volumes of proteinase K ≥ 10 µg/ml in 1% SDS, EDTA, Tris buffer pH 9 at 22°, 0.25 M NaCl then incubating at 50°C for an hour. Wash out completely with buffered salt solution. Destroy any residual proteinase K by running 1.5 vol 0.05% DEPC (diethyl pyrocarbonate) in 0.2% NaH2PO4. Store in 20% EtOH. This has rescued my columns (and others') several times. Manufacturer-suggested extensive washing protocols failed to remove clogging debris.Following
- Ariane Eberhardt added an answer:2How to get rid of SDS from chromatin samples after sonication?
Hi, I am struggling with the problem how to get rid of SDS from my chromatin sample prior to perform ChIP.
My sample is to much diltuted with a buffer containing 1% SDS. As it is necessary to dilute SDS to 0.08% for IP, I have to make a really high dilution of my chromatin sample and the volume of ChIP reaction would be bigger then 5ml. I would like to minimalize the volume of ChIP reaction- I think 0.5-1ml would be perfect. I thought about chromatin precipitation with ethanol, salt and glycogen, hoever i am afraid it will not help as SDS precipitate in low temperature and I am afraid it would bind to proteins in fixed chromatin.
I would be most gratefull for any help!
Hey, can you share the protocol for your Lysis/Sonication buffer? Since I'm using a kit, I cannot tell you the concentration of SDS in my buffer, but I'm diluting it only 5 times prior to the immunoprecipitation and I never had any problem. Can you maybe use a buffer for sonication which is not containing a high amount of SDS?Following
- Anna Konovalova added an answer:5Can anybody explain why when I run a protein with a molecular weight of 9kDa on SDS-PAGE gel it appears as a slightly bigger protein (11kDa) ?
The add Laemmli sample buffer but didn't add BME and didn't boil the sample
Traditional Tris-Glycine protein gel system has poor resolution of small proteins, because free SDS micelles run at the low molecular weight end of the separation "clogging" smaller proteins/peptides and preventing their separation from this front into discrete bands. For proteins < 20 kDa, try running Tricine gels instead. Add a reducing agent as discussed above. You may also add up to 8 M urea, it prevents peptide refolding and keeps bands "sharp".Following
- Jeremy Brooks added an answer:3Why my protein does not elute completely?
Hello everyone, Can anyone please resolve my problem. I am using NTA resin with some preferential metal for understanding the pulldown of proteins by IMAC technique. I elute my protein by using 100mM EDTA. The problem is that "the amount of protein which I get from elution or from protein through is lesser than the amount which I load onto the column". I don't know how I can elute the protein completely?
I agree with the above answers, but also add that if you are washing the NTA column/resin over a number of steps that you collect all flow-through to check it has not been prematurely eluted
You can also check the flow-through when you wash the column after your elution step
Just monitor protein a280 for indications of lossFollowing
- Diaa Ali Al-Raawi added an answer:5What is the difference between MG132 proteasome inhibitor and protease inhibitor cocktail tablets?
I could not detect my target endogenous protein but I can detect over-expressed protein. I used Protease Inhibitor Cocktail Tablets. My friend told me that my target protein could have a high turn over rate so I could not detect the endogenous protein. He recommended me to use MG132 proteasome inhibitor.
1.Should I use both MG132 proteasome inhibitor and Protease Inhibitor Cocktail Tablets to do westernblot or I just use one of both every time I do western blot?
2.Protease Inhibitor Cocktail Tablets and MG132 proteasome inhibitor are used to inhibit protein degradation in western blot. Which one is better?
I am wondering in case if i treated cells with MG123 inhibitor instead added it directly to lysis buffer, this will inhibit cellular proteasome leading to stop degrade protein. if this is natural cellular mechanism, this will affect on endogenous protein level. I wanted to understand because I have noted another strong band (not in right size of my protein) and i want to know if this is band is cleaved product. So I am trying to use different inhibitors, MG123 is one of them. ThanksFollowing
- Zahra Falahati added an answer:7How can I dissolve PMSF (Protease inhibitor) in lysis buffer?
I was trying to dissolve PMSF in lysis buffer. But due to little water solubility it couldn't get dissolve. Which solvent I should use to dissolve it? I will use this lysis buffer for the protein extraction for WB. I am wondering if ethanol or isopropyl alcohol will interfere with protein extraction or their stability. Please suggest. Thanks chandrashekhar.
you can solve it in isopropanol or ethanol also. because pmsf is instable don't add it in your whole lysis buffer stock. you should make it just before you want to add it in your lysis buffer.so it is better to prepare a 200 or 100 mM stock of pmsf and then use it in 0.1-1mM in final volume of your lysis buffer .Following
- Dominique Liger added an answer:4Is it possible to renature any protein after denaturing by removing the denaturing agent?
In theory it is possible but experimentally problematic in most cases. The main issue is to find the best condition to favor the renaturation rather than aggregation.Following
- Maria del Pilar Corena-McLeod added an answer:2Why is my protein not complexing anymore?
I have been purifying a protein about 150KDa and post purification, running on a Native PAGE gel or on SEC Trace, it would elute around 750KDa. In more recent purifications, no complexes have formed and all the protein runs/elutes around 150KDa, as a monomer.
Purification methods are the same, buffers are the same. Perhaps the only subtle differences are Insect cell line, infection efficiency (we use baculovirus), FLAG beads used for purification.
Any insight into why this is happening?
It is not uncommon to see monomers from complexes as large as 750 kDa. Those are very large complexes that are extremely unstable. You could use the same exact conditions and then shake your protein solution with the pipette, up and down and break those complexes.Following
- 7How to stop protein precipitation from pH change in Hi5 cell culture?
We have been using the Baculovirus system for protein expression. When we purify protein from Hi5s, we have to change the pH from about 6.3 or less to 8.0 for purification. When we change the pH some of the protein precipitates out of solution so we cannot purify it. does anyone have any advice to help stop this? We have tried changing the pH of the Hi5 media, which just makes the cells unhappy and take forever to grow. Anything would be helpful!
So it is AFTER purification that the precipitation ensues but does not occur in the conditioned media if left neat? The conditions used for the IMAC is allowing for complete product capture and 100% product recovery, correct? If true then the question in effect is how to stabilize the post IMAC pool with an appropriate formulation buffer which can be quite a matrix using a partial factorial design. Hopefully there's a cell-based functional assay or at least a binding assay or stability indicating assay.
Knowing what is in the precipitate will help tremendously to answer the questions. Likely you can get the precipitate to run on SDS-PAGE if it can be suspended in some water and then a sample of that added to the Laemlli sample buffer with heating. The sample prep process will likely dissolve the precipitate; if not then there is something very much incorrect or dramatically different about the material (co-factors, metal/salts/inorganics, polymerization, lipids, hydrophobicity, etc). In this analysis it will be very important to run the sample reduced versus non-reduced and, if possible, to run the reduced sample after complete alkylation with IAA or NEM. One area to focus upon is any streaking and smearing in the higher MW areas compared to the reduced lanes and assuming the gel wells are not overloaded. You might want to throw away the precipitate but it is not useless.Following
- Boya Venkatesh added an answer:7Can any one help me in purification of my protein?
I'm working with protein purification, the protein has a theoretical pI of 5.95 along with TRX tag. can any one suggest me which buffer has to use for maximum protein elution with Ni-NTA coulmn.? If possible send the reference article links.
thank you Wenjiang Ma for your suggestion. I will definitelu work on this. thank you once again.Following
- Andreia Ribeiro Albuquerque added an answer:3LEXSY L. tarentolae growth problems. tips?
Hi! Anyone working with L. tarentolae LEXSY protein expression system? Despite having used the protocol described by Jena I am having trouble on parasite growth. Any further tips you might have?
Hi all and sorry for the delay in my answer, The growth is the same if you transfect with control plasmid (pLEXSY) or the pLEXSY with my protein of interest. According to the protocol within 5-7 days after transfection you should find colonies of about 1-2 mm and I never found such colony size.
I used both methods of plating with and without nitrocellulose and I only saw increased colony size after inducing the colonies grown in the nitrocellulose membrane in a new plate with Tet.
I am trying to see if I can recover the clones from these induced colonies removing the Tet. and indeed they grew, however I don´t know if the fact that I induced once in the plate might influence future inductions, as I tried to induct a 1:10 dilution in 1 mL and saw no color change after 2 days (even though the colony on the nitro. membrane was pink-ish).
- Ewa Marcinkowska added an answer:3Why do I get smear on SDS-PAGE?
I am working on snake venom proteins and wanted to resolve it on SDS PAGE. I have attached image of my gel. I am getting a smear. Is there any way to resolve it better?
Try to load less protein, overload may be the cause. But in general, gel looks good.Following
- Uma Suganya K.S. added an answer:20How long can I store a SDS-PAGE loading sample (protein) after it's being mixed with dye and denatured (boiled for 5 min)?
How long could the sample buffer be stored? Or should it be prepared fresh all time?Following
- David Morgenstern added an answer:1Which gel to use for LC-MS protein separation?
I did the IP for the separation of specific protein (p53) form the total protein. Now I have antibodies with my specific protein. To remove antibodies and get my protein I have to do gel electrophoresis. So just want to know should I use SDS or native gel?
use SDS PAGE, without any reductant - no beta-mercaproethanol or DTT. this will leave the antibody intact at 150kDa.
- Phanindranath Regur added an answer:4Can we tag our protein with some dye such that we can avoid staining and destaining in an SDS gel i mean some sort of prestaining of our protein?
Can we tag our protein with some dye such that we can avoid staining and destaining in an SDS gel i mean some sort of prestaining of our protein?
Thank you for the input.Following
- Farideh Zarei added an answer:3Does anyone have an efficient protocol for protein extraction from paraffin-embedded tissues ?
I am trying to extract protein from formalin fixed and paffin-embedded tumor tissues. The objective is then to perform western blotting of an membrane protein. Do you have any special recommendation for extracting the protein from paraffin-embedded tissues ? Thanks very much.
HI dear friend
I hope the following articles be useful for your study.Following
- Naman Shah added an answer:6Any solution for problems with Batch Method Purification of his-tagged-proteins?
I have to purify a protein with 6xHis tag, but at the time of dissociating the protein from the resin (Ni-NTA) all protein remains in the resin. A feature of my protein is that it is rich in histidine in some sectors. I increased the amount of up to 500 mM of imidazole but I have not had good results. In advance thanks for the help.
You can also try to engineer a TEV site between your his-tag and protein. So after it gets bound to Ni-NTA, you can cleave the 6-his off the protein. Granted that your histidine amino acids of the protein will still have some affinity. You can then try eluting with imidazole.Following
- Dan Weaver added an answer:4Akta Pure, any resources for IgG purification methods?
Much of my experience for IgG purification is in UNICORN 5.0 on an Akta Explorer and I'm looking for resources in setting up pH gradient methods for elution in UNICORN 6.4 on an Akta Pure.
Does anyone have any general methods available?
This was work I did between 2009 - 2011 and I don't have access to all the details any longer - especially the script's code - property of a different company. This code is not difficult to write - the language structure is a form "script" which is the same for the BIAcore/Akta systems. Depending upon the features you want - the basic code should take only a couple of hours to write and debug. Additional features are added along the way to improve the process - means improving your life-style around the process. Slow Flow rates improves your resolution to an extent - yet the value I suggested works quite well - a good comprise. These are standard columns sizes (geometry's) which are not very difficult to scale up - all well designed by the manufacturer. Time to Validate the working code is dependent upon pushing through a QA system.
All my antibody's looked the same as far as resolution (see below) - I did not have culture supe's or bioreactor media to purify from - to much volume to ship across country - too expensive.
We had a mixture of IgG1 and IgG2 in that library (40 - 45 Ab I prescreened by BIAcore for functional activity) - probably more IgG2's than IgG1's. The vendor scaled up slightly - purified using the classical method you described - froze them then shipped them to me. I biotinylated with a Chromogenic Biotin (Solulink). Once Ab-bio linked to streptavidin latex beads, these latex beads aggregated - bad news for lateral flow assay. I used the pH linear gradient before biotinylation to solve the aggregation problem at that later - assay development back on track - worked like a champ! I had about 2-4 mg of antibody to pull this off to demonstrate proof of concept for the assay design.
Citrate was not a problem in our processes - I went quickly to the biotinylation (raise pH with Borate - Amine chemistry NH ester). The product was loaded onto a 120 mL Sephacryl column (~0.25 mL sample @ ~10-15 mg/mL, ) and purified by gel-filtration to purify. Keep in mind that the gel-filtration process was in place and used even with the frozen Ab - so the linear pH gradient solved an issue in our processes. To this day I do not know what problem I solved - I just could not repeat the method that my vendor used to solve this problem - I needed a different result from my vendor!!!
Sorry I didn't have the code to send off to you - If you want help with the code send it off to me - however - I want to reassure you that it is pretty easy to write and debug.
looking forward to here from you!
- Kambiz Gilany added an answer:4How many 2DE gels run for clinical proteomics trial?
Hej Everybody. I am looking for to get answer how many 2DE gels I have to run for a pilot study in clinical proteomics? Should I pooled the sample and run three 2DE-gels or just run 3 good 2DE-gels and run western-blot on the potential biomarker? How high the fold changes should be for quantification to be real? I read different paper and some reported more than 1.5 fold change etc. The best value is not 2 fold changes?
TX everybody. Can anybody refer me with reference about 3 run samples and 3 2DE gels?
- Michał Szlis added an answer:2How to measure the concentration of 43RFamide in sheep CSF?
Is there any ready and checked ELISA/EIA/RIA kits or maybe I should think about chromatography assay?
thanks Simon, this is a very good idea about which I forget =] I need definitely to try it.Following
- Imdad Khan added an answer:8On western blot, how can i eliminate the non specific reaction in the form of black dots which represent the binding of first antibody to skim milk ?
i am doing western blot for my research study and i am getting the black spots on the membrane after developing which indicate the binding of first anti-body to skim milk. how can i eliminate this problem.lot of thanks for the sincere help and guidance, i will certainly consider the suggestions offered by you peoples.Following
- Mark K Chee added an answer:3What is the function of methanol an acetic acid glassial in Staining solution and Destaining solution in SDS-PAGE?
Some site said that methanol will shrink the gel. Some site said acetic acid glassial fixed the protein, but i don't know their special role in each of the solution since the two solutions have methanol and acetic acid glassial.
When I read how commassie blue work with protein there are low pH2 and above pH 2. How commassie blue work? Is acetic acid glassial has corelation with the pH in staining solution?
However, I think this article gives a lot of food for thought about the lore behind CBB staining: http://www.labtimes.org/labtimes/issues/lt2008/lt06/lt_2008_06_53_53.pdf
I hope that helps!Following
- Boya Venkatesh added an answer:2Need protocol . if any one have please share ?
I need a protocol for purification of thioredoxine fused protein with Ni-NTA column from bacterial system.
if any one have the protocol please share.
thank you for your reply sneha madamFollowing
- Rachel Bell added an answer:3Optimising a PCR reaction where I am currently getting no band?
I am in the process of preparing to do a RACE-PCR experiment and am testing some primers for this. I have designed one forward primer and two reverse primers (these will be used in a nested approach in the RACE-PCR). When I ran the PCR reaction today to check fragment size amplification on plasmid DNA containing the gene of interest, one forward/reverse mix gave a strong band as expected (~130bp), whereas the other mix did not give the expected band (at ~180bp).
These primers look good on Primer Blast (good specificity to the target, low self complementarity etc.) with similar Tms of ~64C.
The conditions I used for the experiment were as follows:
95C for 2 min
25 cycles x 95C for 30s, 60C for 30s, 72C for 30s
72C for 2 min
I am still confused why I am getting such strong amplification with one primer set and no band whatsoever with the primer stock. Could anyone give any suggestion to improving my cycling protocol or any other points to consider? Something that may or may not be relevant is that I do get a fairly strong primer-dimer band at the bottom of each lane, I am probably including too much primer into the mix, but this is something that is consistent between the successful and unsuccessful reaction.
Hi all, I have tried adding DMSO with no luck, and will try the gradient PCR today.. if no success there I will design a new primer. Thanks for your help!Following
- Ralph Rozier added an answer:40Does anyone have any experience running ERIC-PCR?
I have run ERIC-PCR a number of times and have failed to get bands in the manner best depicted in peer reviewed journal articles.
I am running it for Salmonella using the following primers:
ERIC1R: 5’-ATG TAA GCT CCT GGG GAT TCA C-3’
ERIC2: 5’-AAG TAA GTG ACT GGG GTG AGC G-3’
I have run over 20 different runs altering as many issues as possible each time.
I have changed the amount of template DNA (10ng to 1000ng), the amount of Taq, the amount of MgCl2, and the amount of each primer (10 mM to 100 mM).
I have run the gel at 100v for one hour and for four hours.
I have run the gel at 40v for four hours and seven hours.
I have run the gel at 85v for six hours and twelve hours.
My gel is 1.5% agarose with 1x TAE. I mix in ethidium bromide before running.
My New England Biolabs Taq, Buffer, and dNTPs work fine as I have run it with another PCR protocol.
I have run the PCR at four different protocols that last anywhere from one to six hours.
I got some bands with a short protocol, but they were only on a short gel ran at 100v. They looked nothing like the papers do.
I have used both Salmonella from a known sample and an environmental sample. I have used boil lysis and Qiagen kits to extra DNA.
I am getting nowhere.
Thanks! I will try.Following
- 9I am trying to express and purify a protein that belongs to lipid binding protein family with reference to the actual crystal paper?
Though i followed the expression and purification conditions exactly in the paper, i could not see the expression. What could be the reason? Did you face same problem in reproducing the results of protein purification?My vector is pET28a expressed in BL21(DE) at 0.4mm IPTG at 16C for 20 hours. The sequence is in frame.
Seems you might still have culturing concerns, not just for expression level but possibly for levels of degradative activities. If this is a membrane associated protein and the membrane domain is still present then a significant portion of product may be in the pellet from the homogenization with some partitioned into membrane vessicles/fragments in the supernatant. If membrane association is possible and can obscure the tag then the effect of DDM detergent may be interesting. In any case, for crystallization goals, you'll likely need protein levels that don't require Westerns for detection other than for identification. Since scale up is usually not linear, then there's the likely risk that a greater amount of fermentation will not produce more total product.Following
- Ki Oh added an answer:4How important is lysis buffer volume for purification?
I am expressing and purifying recombinant GFP/6His protein from Sf9 cells. Recently after what looked like a solid expression (green GFP pellet) from a 500 mL (2e6cells/mL) culture, I resuspended the pellet in 10 mL of ice cold lysis buffer and froze them as pearls in liquid nitrogen.
The 10 mL was purely empirical but I have seen protocols where it suggest 4mL lysis buffer / 1~2e7 cells. Meaning I would've had to use 200mL of lysis buffer.
I know other protocols go by pellet weight but in the end, what is the downside of having a concentrated lysate? Poor lysis? Sub-optimal purification?
Would it be recommended to dilute the frozen pellets with more lysis buffer before sonicating and purifying?
Thanks in advance
Thank you for the answers!Following
- Gary Laco added an answer:5What is the cause of uv-baseline negative drift ?
In our process for purifying some proteins, we should use 2-me (beta mercaptoethanol) at concentration of 4% w/w in elution buffer.
and we have a serious problem with uv-baseline. it drifts negatively.
We know that, this problem comes from 2-me. Because when we use buffers without 2-me, it's all good.
but why there's a negative drift in baseline with 2-me? and how can we fix this?
Our columns are: DMAE Fractogel and TMAE Hicap Fractogel.
equilibration buffer is 20mM Tris + 4% 2-me w/w
and buffer B is 20mM Tris + 500mM NaCl + 4% 2-me w/w
we know that concentration of 2-me in our buffers is too high (~700mM) but we should use this concentration of 2-me to prevent aggregation of this protein.
We greatly appreciate if someone try to help us.
Why not use a different reducing agent like TCEP?, it is more expensive by is not volatile like BME so you would avoid the fumes/odor and it is a better thiol reducer on a molar basis so you would not need nearly as much, plus other advantages, see: https://en.wikipedia.org/wiki/TCEPFollowing
- 9Why mbp-cleaved protein disappears after anionic exchange?
I am purifying a protein (47 kDa) that is fused onto an MBP (42 kDa). The PIs for both proteins are 6.48 and 4.9 respectively. After purifying it using a dextrose column, I have buckets of fused proteins (up to 30 mg/ml). I dialysed my proteins into a reaction buffer to rid of the maltose and to optimise the cleavage of the tag (20 mM Tris, 50 mM NaCl and 1 mM CaCl2 pH 7.5). Consequently, I tried cleaving it at small scale (10mg of protein with 100ug of TEV protease with 0.005% SDS) for 6 hours.
After cleavage, I dialysed my samples into a buffer (20 mM Tris, 50 mM NaCl and 1 mM CaCl2 pH 6.4) for anionic exchange. Theoretically, my proteins should be neutral and only other proteins would bind onto the anionic exchanger. The picture of the gel shows the flowthrough and the elution of my proteins. Lanes 1-3 show the elution of my protein while lanes 4-9 shows the elution of other proteins. So far, the bands corresponds to the size of my protein. However, whan I mass-spec my the bands on this gel, the results show that the bands on lane 3 is mainly MBP. the concentration of the flowthrough is 1.63 mg/ml.
This is my first time using MBP and purifying this protein. I can't think of where my protein might have disappeared. There were no signs of precipitation in the solution. Any ideas?
In the future I will be optimising the expression of my protein. e.g. inducing at room temperature overnight, adding glycerol in my buffers and conducting my experiments at 4C. I would also run my samples through another dextrose column after cleavage to rid of the MBP although I am not sure as to how this would improve my technique and lessen the loss of my proteins..
This is what I suspected could happen (yet, still no proof) and why I suggested running your gel samples both reduced and non-reduced as well as running a sample of the column strip (material balance). A column strip is not a high dextrose or high salt elution but to run something like 6M guanidine HCl or urea with maybe 0.5 M NaOH to be sure all protein is removed from the resin. Would you be surprised if those higher order bands collapsed to monomer mass upon reduction?
Also it seems the monomer may not be recovering well but you don't know until you do material balance which will imply a lot about what could be occurring. Covalent aggregates, if you prove their presence with reduced gels, indicate that the target moiety of the fusion is not folding well so that both aggregates and monomer may be mis-folded, albeit the MBP may be fine. Mis-folding can definitely obscure your TEV site; no cleavage with the site buried . Have you done a basic experiment of incubating the pre-dextrose column fusion molecule with maybe a 1:25 ratio of protease in solution (not on-column) and then running the gel reduced vs non-reduced (you might also pick up the condition of the protease)? I assume this was an E. coli expression system and so no glycosylation? I assume this is a eukaryotic protein with cysteines?
Another possibility: Etch virus protease is a cysteine protease but is there any trace of reducing agent and EDTA in your cleavage reactions (didn't see in your description)? TEV protease can also auto-inactivate but I believe there are versions that don't do this. If your protein needs to form disulfides for correct folding then you have to consider the order of folding in conjunction with a reductive cleavage condition. Run the process reduced and then fold? If is folds firstly with disulfide formation and then requires partially reductive cleavage conditions then what might happen?Following