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Protein Biochemistry - Science topic

Protein biochemistry is dedicated to the study of the chemical and physical structure of proteins, as well as complex chains of amino acids.
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I would like to use 100% methanol (-20 degree C) for fixing monolayer cell culture for ICC-type of procedures. Is there any requirement on the grade of methanol to be used? E.g. Sigma has this (Cat. No. 494437) methanol "Suitable for protein sequencing, BioReagent" and this (Cat. No. M1775) listed under "fixatives", while we have analytical grade methanol (for preparing Western blot transfer buffer) in the lab. Can someone advice which grade of methanol should I use? Many thanks!
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Saravana, it seems that fixing cells with -20 deg C methanol is recommended by more than one source.
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Most people say that imidazole doesnt affect the majority of downstream applications for purified proteins but it is a chelator, so you would think that it might chelate the Mg2+ in solution and inhibit Mg2+ dependent reactions. Anyone seen any papers that discuss what concentration Imidazole will chelate magnesium or manganese ions in solution?
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Not only ligand concentration but pH is also an important factor of chelation .
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I performed protein precipitation using 40% ammonium sulphate precipitation as follows
1. 5.5 grams of ammonium sulphate salt added very slowly over a 30minutes time with gentle stirring for a 25ml of protein solution. (40% saturation) on ice
2. The solution is kept on ice with stirring for 6  hours.
3. The precipitate is pelleted down with ultracentrifuge. 
4. The SDS gel of the pellet shows strong protein band at the expected position (37kDa)
I then increase the buffer quantity to 100ml which is more than what I had initially started with (25ml) which means the salt concentration is way less than 40%.
Upon dilution it dissolves i.e. I can pass the solution through a 0.22micron filter without problem which means that the protein is no more a precipitate in the solution ...but when I try to concentrate the solution using a 10kDa filter after 20 minutes of spinning at 4000rpm the solution in the centricon starts showing precipitates....which means the salt is simply not moving out of the centricon.
Even after several washes (5 to 6 washes)the protein simply seems to remain as precipitate in the centricon.
what does this mean?  What else can I do to remove the salt from my protein solution?
Thank you very much for your time
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You can remove ammonium sulphate from protein sample by dialyzing the protein against low salt concentration, then dialyze
with buffer with no salt. To check that ammonium sulphate has been completely removed from protein solution present in dialysis bag .Take 1 ml of solution( from container in which you have put dialysis bag ) in test tube and add pinch of barium chloride .A white precipitate is formed which is dissolved by adding few drops of nitric acid which means ammonium sulphate has been completely removed from dialysis bag otherwise keep on changing buffer solution of container with fresh buffer containing no ammonium sulphate. This will take at least 2 days to remove ammonium sulphate. After that you can concentrate the protein in dialysis bag . For this a simple solution is to put the dialysis bag in front of fan for several hours or even overnight provided that room temperature is not warm ( to prevent protein from inactivation or aggregation) .
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From transient e get very high levels of secreted recombinant protein with a N-terminal his-tag, but our purification results are poor (less than 5% yield). We have tried desalting the supernatant, using 5ml or 1ml HisTrap FF or HisTrap excel columns, with or without Imidazole in the loading buffer, but so far there is hardly any effect.
Expression and purification is determined by another tag (c-terminal) on the protein which we quantify on a bead-based assay.
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Expi(CHO or 293) media contain EDTA, dialyse or use chelator stable resins
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Hello. I work with bacterial spores. Currently, I am reading an old project report of a student who performed protein digestion from spores and estimated the amount of dityrosine (dT) present in his sample. I will first give some details and then ask my question.
(A) It is suggested that in the outermost protein layer of spores there are dityrosine (dT) cross-links present and it is estimated that every 60kDa protein from the spore outer layer has 3 dityrosines present (Pandey and Aronson 1979; Properties of the Bacillus subtilis Spore Coat).
(B) The student's (who's report I am reading) estimation was based on fluorescence measurements. dT can be detected at pH 2 with ex. 283 nm and em. 410 nm or at pH 9.5 with ex. 315 nm and em. 422 nm. For this measurement he used in vitro synthesized dT (Malencik et al., 1996; Dityrosine: Preparation, Isolation, and Analysis) with absorbance of 1.45 and ε (318 nm) = 8.7 mM-1· cm-1. From these parameters it was estimated that ~0.167mM dT was synthesized and it was used as standard for fluorescence measurement. The theoretical maximum of enzyme-catalyzed (in vitro) conversion of tyrosine to dityrosine with the above described method is ~ 25 % i.e. 0.25 mM or 250 nmol in 1 ml.
(C) The extracted protein amount that was used by the student for dT estimation was approx. 78-100 ug.
(D) The student mentions that "based on the fact that 60kDa has 3 dT, the fluorescence of 100x diluted dT standard would be comparable with that of the 2x diluted protein extract sample".
My question is, can someone explain to me how point (D) is valid?
I am attaching the papers and the extract of student's report. I hope some one can help me.
Greetings,
Wishwas.
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Typical positive congo red spectral assay for binding to amyloids would be an observation of spectral shift from 498nm (CR only) to 540nm (in presence of amyloids). However I've tried the both the spectral assay and the birefringence assay (http://www.assay-protocol.com/biochemistry/protein-fibrils/the-congo-red-birefringence-assay) with an amyloid sequence AB(27-32) peptide that forms amyloid fibers, the spectra I've got does not shift to 540nm, there was only an increase in absorbance at 498nm. Under polarized light microscopy the fibers do appear to be apple green and birefringent after staining with Congo Red though. What is wrong? Or is it normal to get an increase in intensity of the spectra instead of spectral shift?
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Also had such problem. I use Congo Red for studying amyloids in a biofilm matrix. With comparable visualization of the staining density, I found that 540 nm works best for me, no more, no less.
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Hi,
I am studying a simultaneous proton transfer, bond breakage and nucleophilic attack (by water molecule), using US approach for which I had already performed 5ns QM/MM simulation.
All three reactions takes places in a single step (Inversion mechanism for Glycoside hydrolase). Now, I am confused in defining the restraint variables.
I have selected 4 Reaction Coordinates:
1. RC1: Proton transfer from Base residue to leaving group
OE1-HE1 -> C----O4 (this glycosidic bond breaks and HE1 is transferred to O4 )
So, the reaction coordinate for this reaction is difference in distance between OE1-HE1 and O4--HE1.
2. RC2: Glycosidic bond breakage:
C-----O4 -> C O4. The reaction coordinate for this reaction is the distance between C and O4
3. RC3: Nucleophilic attack by water:
H(i)O(w)H(w) [this is nucleophilic water] ---- C (anomeric carbon of the broken glycosidic bond). The reaction coordinate for this reaction is the distance between C and O(w).
4. RC4: Proton transfer from water (H(i)) to Acid Residue
H(i)O(w)H(w) -- OD1 (Acid residue). The reaction coordinate for this step is difference in distance between O(w)-H(i) and OD1-H(i).
For the RC2, I have made the following restraint file:
# distance restraint
&rst iat=8122,8132 r1=0, r2=1.8, r3=1.8, r4=5, rstwt=1,-1, rk2 = 500.0, rk3 = 500.0, /
I have increased the the value for r2 & r3 by 0.2 and upto 3.4. I am not able to understand what should be the value for r1 and r4 ? Could anyone pls comment on it and explain it briefly?
I also not able to understand how to make the restraint file for difference in distances between two set of atoms, as in case of RC4 and RC1. I would be helpful for me if somebody explains it too with an example.
I also want to visualize all the four reaction steps so which trajectory files from all the four RCs I should see?
Since I am new to US, it would be a great help if somebody can guide me through this.
Regards
BHARAT
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Bharat Gupta how you become able to write your umbrella sampling input disang file (as mentioned in the amber tutorial) using the LCOD method. Can you tell me, please?
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Hi, I am running a size exclusion chromatography experiment with a buffer containing Potassium Acetate as a salt. I analyse these fractions through SDS-PAGE. After boiling my SEC fractions in laemmli buffer and leaving to cool to RT I notice a loose precipitate in the tube. After spinning this down and loading the supernatant on an SDS-PAGE gel I can visualize my proteins of interest.
I was wondering if anyone could help me understand what this precitipiate is? I believe it could be potassium dodecyl sulphate? Is it expected that this would also precipitate some of my protein and reduce my soluble yield that I can visualize by SDS-PAGE?
Thank you.
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You are right. Potassium dodecyl sulfate is much less soluble than sodium dodecyl sulfate.
potassium DS 0.415 mg/ml.
sodium DS 200 mg/ml
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Based on this paper
,TEV protease can cleave between the Gln and several amino-acids (besides Gly/Ser) with acceptable efficiency in its recognition site.
Therefore, it's practically possible to purify many proteins (without an extra residue at the N-terminal end), by using affinity chromatography.
I was wondering if anyone could share their experience/knowledge using TEV protease to cleave between Gln and Met?
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Sorry, I have no experience with the specific case of Q/M cleavage, but remember that most proteins whose second residue is a small sidechain aminoacid actually have their N-terminal methionine removed by MAP while still being elongated in the ribosome.
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Good afternoon,
I try to extract cell wall proteins from green macroalgae, I have tried different protocols and one of them included step of EGTA extraction, however this fraction is jelly like, so I expect polysaccharides from cell wall to be present. When loaded to SDS-PAGE, it was obvious that the gel pores were blocked and no bends were obtained, just light blue streak. According to Bradford, in this fraction I have quite a lot of proteins with which I would like to do Western Blot... I have already tried aceton precipitation, but still a lot of polysaccharides remained....So please do you have any recommendations how to get rid of these polysaccharides?
Thank you very much,
Tereza
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Hi everybody,
we have a protein which is toxic if it is highly expressed (SV40 promoter). Therefor I had the idea to use a minimal promoter like the minimal CMV ( GTCGAGGTAGGCGTGTACGGTGGGAGGCCTATATAAGCAGAGCTCGTTTAGTGAACCGTCAGATCGCCTGGAG ) to get only low amounts of the protein. However I found several "versions" of the minimal CMV and would like to have the one with the highest basal expression. Can somebody recommend me on and send me the sequence. Iam also happy to get other suggestions for promoter with low basal expression in a mammalian system. 
Thanks a lot,
Flo
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Dear researcher, minimal promoter refers to a minimal sequence of a native promoter (mostly core promoter) that could express a downstream gene. So, that not mean minimal promoters' expression is low or in minimal level. CMV and SV40 are virus derived promoters and they are so strong. However some promoters are engineered so they have lower levels of expression compared to native promoters, like engineered ADH1 promoter. Good Luck
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Most of the researchers working on protease inhibitory assay, they cited the Oyedepo etal, 1995, but I couldn't get the original paper. I followed their protocol, replicated 11 times, but unable to get the readings as absorbance was read very high. Even the Aspirin and Diclofenac sodium at the dose of 10-100 microgram/100 microlitre as well as 1-100 microgram/ml could not give the reading . I am wondering why I can not get the readings even on standard drug following their reported protocol. Does Trypsin (0.06 mg) have any specific concentration in their protocol ?
Oyedepo OO, Femurewa AJ (1995). Anti-protease and membrane
stabilizing activities of extracts of Fagra zanthoxiloides, Olax
subscorpioides and Tetrapleura tetraptera. Int. J. Pharmacog., 33: 65-69.
Please provide any info on the protocol of this paper and suggest another protocol for protease inhibitory assay if possible.
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OK
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For comparing pure and adulterated honey
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AOAC 998.12
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I’m working with a rather complicated ping pong, bi-bi double displacement reductase enzyme that oxidizes NADH and then reduces a substrate. Interestingly, the enzyme has intrinsic NADH oxidase activity in the absence of the substrate. Of course, upon addition of substrate, the velocity of the reaction increases and NADH is consumed more rapidly. The intrinsic activity introduces a baseline velocity and is complicating our kinetic analyses a bit. Complicating it even further is a noted double substrate inhibition pattern (apparently not uncommon with ping-pong mechanism enzymes). We’ve run a full course of analyses, varying the NADH as well as the substrate and we're now trying to fit the data to an appropriate equation. My question is: How do I take the ‘intrinsic activity’ of the enzyme into account (if I need to at all)? Can anyone recommend an equation and appropriate software that can do this? We currently have SigmaPlot, GraphPad, and Enzfitter (for the double substrate inhibition model). Thanks in advance for any guidance, ideas, and/or discussion.
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قياس فعالية الانزيم في الجزء المراد دراسته ومن ثم اخذ برنامج احصائي هو spss
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S.coelicolor produces red and blue antibiotics, which has affected the use of Bradford method to measure protein concentration of cell extract. The most commonly used methods are based on colorimetry which I think can be affected by pigmented antibiotics. Are there any better methods? I have also thought of using internal reference. What kinds of internal references are used?
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Hi! I have the same problem and I wonder if you managed to solved it? Regards. Nancy
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I have expressed my 70 kD protein (pI 6.7) in BL21 E.coli cells and purified it using TALON metal affinity resin (cobalt-based resin). My elution buffer is 20 mM Hepes Ph 7.6, 500 mM NaCl, 10 % glycerol, 1mM TCEP, and 150 mM imidazole. My previous experience with this protein had shown that it tends to form some sort of dimers/aggregates thus I added 1 % octyl-glucoside (OG) and 5 mM EDTA to the eluted protein that were in a buffer mentioned above. Although addition of 1% OG was borne fine, upon addition of 5 mM EDTA the solution went cloudy and after 48 hour incubation at 4 degrees there were lots of aggregates in the solution. I spun down the aggregates and tried to resuspend in 8M urea but it wasn’t successful attempt.
I thought addition of EDTA might be beneficial through chelation of any leached cobalt ions. Is it a possibility that chelating leached cobalt ions also precipitated my protein?
Or chelation of useful divalent ions knocked my proteins out and precipitated them?
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I know this question is quite old already, but since I found It as the top results on my google search I thought I might help any future visitors.
It is true that EDTA can cause problems for his tagged proteins, the stability of a quality Ni-NTA agarose resin is about 1 mM here. Every concentration above that will have severse consequences. When working with NTA or IDA based resins at least.
The INDIGO ligand has an EDTA stability of 20mM, so 20 times the concentration of quality Ni-NTA. That should be used here.
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I am working on a human helicase protein and when I load them on a SEC column Superdex 200 (GE), I don't get a good recovery. In addition, when I load my protein-DNA complex into the same column the complex does not come out at all. Had anyone experienced a similar problem?
Also, had anyone here tried to purify a protein-Holliday junction complex? I have a mixture of two oligomers (homo) in my protein, and one of the oligomer binds to HJ, while the other does not. For further studies I would like to purify the protein-HJ complex from the non-binding oligomer of the protein, but not finding a good way. Please share your thoughts/experiences.
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hi, i'm experiencing something similar. there is a protein portion that elute during a log range of volumes and at the BN-PAGE analysis it looks quite heavy. i don't understand why it elute along all the volumes from the aggregates range to the monomers range. i was thinking the same, that my protein being highly glycosilated stick to the column. dos it make any sense? any suggestion?
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Hi,
I am doing a pulse chase experiment and looking for 35-S methionine/cysteine incorporation in my protein of interest. Following gel run, I directly dry the gel (10% SDS-PAGE, 1.5mm thick, BioRad mini) in a gel drier for 1 hr at 80C. Then I expose the Xray film with an intensifier screen. My signal intensity is either very weak or no signal at all even after longer exposure (I went up to 72 hrs.). Although, several things might impact the result, however, I was wondering if fixation is necessary after the gel run (before gel drying, I guess)? If so, how would you do that?
I would appreciate any comment/suggestion!
Thanks
Sumit
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Hello, plastic foils or glass plates is necessary for fixation of polyacrylamide gels to the support autoradiography. When using glass plates as support, autoradiography creates problems which can be completely avoided when the autoradiography plates are heated for 30–45 min at 55–60°C prior to gel polymerization.
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I am working with preventing the fibrillation of lysozyme. The samples I'm analyzing contain aggregates that are not fibers and of which I don't know the secondary structure. When using Circular Dichroism I consistently get a strong negative peak at around 230 nm, which is not consistent with alpha helix, beta sheet or random coil. I was wondering if this could pertain to an oligomer of lysozyme fibrillation or to some other kind of aggregate. I appreciate anything that might shed light into this phenomenon!
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First thing is to check is if anything in the sample buffer is absorbing strongly either in this region or at shorter wavelengths.
If the peak is specific to the protein, a positive peak at 230 nm in protein CD spectra may be an indication of a strong pi/cation interaction, usually between a charged amino acid like lysine an and an aromatic amino acid like tryptophan or a metal and an aromatic amino acid.
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I have a system where the protein is embedded in water box and has a non-polar interface consisting of non-polar organic molecules such as DCE. We are trying to study adsorption of protein at the interface under the effect of electric field in z-direction. The protein reaches the interface but has undergoes large conformational changes.  I would like to measure the orientation angle between the helical domain of the protein and the interface (water-DCE). How do I define the normal to the surface/interface and calculate the angle?
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Dear Neha
I prepared some power point presentation about Protein _Ligand Docking Studies.Try to see its give some idea regarding your works.
Kindly see attachments.
Thank you
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protein purification, protein concentration
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Dear Ahmed.
you can wash them with milliq water to remove all the Salt and store at 4°C in 20% of ethanol also for long time. When i was student and we had strong buffer restriction i re-used it several time and it works. however while is simple remove the Salt and other buffer component , it not so simple wash well the inside membrane from the protein and to avoid cross contamination.
i reccomend to you to re-use it only for different preparation of the same protein but change it once you change your protein.
for desalting i suggest to you ti evaluate the pd-10 desalting coloums that are fast and do not stress your protein sample.
you can find some more informations about it at page 3 on my blog: ProteoCool
ProteoCool n°8 (Buffer Exchange Methods overview) and
ProteoCool n°11(Rapid buffer Exchange by PD-10)
ciao
Manuele
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I`m seeking for RFP plasmids and wanted to order pmCherry-C1 from Addgene, but theay are not available due to their distribution via Clonetech. But i found the very similar plasmid mCherry2-C1 with this explanation http://www.addgene.org/static/data/plasmids/54/54563/54563-attachment_mod-3zR-cqRQ.doc
Can i use both plasmids equally? Thanks for any advice or experience!
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Zach Hensel "Also, I've found that in E. coli, mCherry maturation efficiency is very bad at 37 C, but OK at 30 C."
According to the original developers (http://dx.doi.org/10.1371/journal.pone.0171257), mCherry2 was evolved in a similar way as a superfolder GFP - essentially selected for E. coli colony brightness when fused to ferritin. So I would expect it to be better at least in E. coli. Though, in the original publication they say it does not give brighter fluorescence in E. coli but only shows less intracellular toxicity (hence, larger E. coli colonies).
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I use the Bradford method to quantify my protein with the Pierce BCA Kit assay. Particularly I use the microplate assay, 25 uL of a standard solution is mixed with 200 uL of a reagent, and after incubation at 37 C for 30 min, the absorbance at 562 nm is read. There are 9 standard points ranging from 0 - 2000 ug/mL (2 mg/mL).
I got a good equation which was y = 0.001x + 0.0551
R2 = 0.9944
I purified a recombinant protein with affinity chromatography, it's histidine-tagged. I kept all the last eluent.
What I want to ask is,
After concentrated my protein with an amicon centrifugal filter unit, I obtained at least 500 uL.
I readily quantified this solution with the same condition as above i.e. 25 uL aliquot was mixed with 200 uL of the reagent, left for 30 min at 37 C, and A562 was read.
Let s say the absorbance is 1.500, and with the equation I'll get:
1.500 = 0.001x + 0.0551
x= 1444.9 ug/mL or 1,44 mg/mL
Yes, I could use this concentration for my enzymatic assay. But what baffled me is....
How much the actual amount of the protein in my 500 uL ?
Because I want to incorporate this protein into a certain medium. And my teacher said to me, no need to lyophilize my protein. Just directly use it.
But How could I adjust the concentration of the protein? e.g. 0.01%, 0.025%, and 0.05%. The media is a powder. The protein needs to be mixed with the powder and then a homogenous dough is obtained (with the addition with ddH20).
Thank you
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From my experience:
All photometric assays that rely on a standard curve will always just give you a good estimate. Use a different standard (e.g. IgG vs BSA) and you get different values. Still ok for crude mixtures. For purified protein we compared amino acid anaylsis with OD280 based on the sequence-derived extinction coefficient (the theoretical one) and they were close enough to give me piece of mind that OD280 works well for most of our proteins.
Cheers
Peter
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any technique which can be used to tell the functional and bioactive properties of keratin
any other theoretical reason
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I have been analyzing 90-amino-acid fragment of a protein using HSQC and triple-resonance (1H-15N-13C) NMR. I know roughly where arginine, lysine, and tryptophan side-chain peaks lie on the HSQC, but how can I definitely distinguish what is a backbone and what is a side-chain? In addition to the HSQC, I have several 3D experiments (e.g. CBCANH and CACB(CO)NH).
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Arg Nε-Hε and at low pH Arg Nη-Hη and Lys Nζ-Hζ are visible in HSQC but these are aliased/folded peaks. Go for an open Sweep Width (in 15N dimension) HSQC where these peaks will appear at their exact 15N ppm values and you can identify them easily. If you go to the R/K side chains' 15N plane in CBCANH, you will observe CD and CG of R or CE and CD of K. The Trp side-chain usually shifted downfield and appear near the bottom left corner.
Start with the assignment, you will understand which one is the backbone NH because in CBCANH and CACB(CO)NH they will show only the CA and CB (these you have to identify based on their C ppm values). But the side chains will be different.
Best wishes.
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I am wondering if there are any web-based programs, like ExPASy that can "predict" a protein's sequence based on structural input. For example, ExPASy SWISS-MODEL can predict protein structure based on sequence input, but does the vice versa exist, either on ExPASy or elsewhere?
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Depending on what your need is - if you are working with a structure that has been deposited in the PDB, you may want to use the sequence associated with that file in the PDB, which (usually) is the sequence of the construct used - including residues that are not resolved in the structure because of disorder. This sequence is also to be found in the SEQRES records in the header of the PDB file. If you need exactly the sequence represented by the coordinates, the servers listed by
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I am using the "ImageJ" software in order to quantify the protein bands on the SDS-PAGE, but I'm not sure it's the best software for this purpose. Does anyone have the experience of using other software to do this? Is there any professional software for SDS-PAGE densitometry?
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Usually, the imaging instrument comes with software for densitometry. I have used the Molecular Dynamics software and the Azure Biosystems software. They are pretty similar in all the important respects.
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Is there any trustworthy survey/database/reference where I can know the number of protein families that interact with a specific substrate/cofactor?
I would be interested in knowing the number of proteins interacting with:
.FAD
.FMN
.Riboflavin
.TPP (thiamine pyrophosphate)
.NAD(P)H/NAD(P)+
.ATP/ADP/AMP
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For each of the specified molecules there shall be large number of proteins/ enzymes that can bind. For example cholesterol oxidase can bind both FAD and riboflavin as these are cofactors for this enzyme. One must search databases for each of specified molecules. Use bioinformatics tools....
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My protein is in Tris buffer pH-7.5 and has 9 free cysteine. I tried twice but didn't get an absorption at 412 nm.
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Do not use Tris buffer. Use phosphate buffer. I have seen that the TNB absorbance fades when an organic buffer is used. Also, make sure the pH is basic at the end.
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So far, I have been using SMA to lyse my cells as also to solubilize my membrane proteins and purifying them by Ni-NTA affinity purification (since my protein is 6x-His tagged). I am trying to elute the protein at 250 mM imidazole concentration. But somehow it seems I am losing my protein in the process. Can anybody suggest a better approach from their experience? Thanks much in advance!
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Hi Suparna,
Since SMALP is negatively charged, it may interact with cations, i.e. Ni++. Try high salt concentration, i.e. 500mM NaCl, and increase imidazole (up to 500mM). Good luck!
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Hi All,
I have a fusion protein (fused with a N-term His6 Tag) that I want to try the TEV cleavage on (ENLYFQ/C), cleavage is expected to happen between the Q and C. I tried different conditions such as varying concentration and ratios of Protein:TEV, time etc but in all the cases I am only getting 50% (approx) cleavage, the rest of it just doesn't go to reaction completion no matter what you do. I am suspecting that this is probably because of my TEV cleavage site is buried and thus inaccessible to TEV protease and I have indirect proofs of that as well (my protein didn't bind well to IMAC column while in TBS buffer but the binding improved when It was in 6M Guanidinium Hyd solution). I am also using 1mM DTT during my cleavage but that doesn't help much as well. Suggestions and sharing experiences are most welcome.
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haha glad your problem is solved. it would probably help if you could share how you optimized your conditions for anyone else visiting this thread with a similar issue in the future.
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Curious to find out.
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You can always calculate the pKa(s) with quantum chemistry method. Here is an improved model for your reference,
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I'm designing an assay to test for release of pyrophosphate due to dihydropteroate synthase-like enzyme activity using malachite green reagents. One of my substrates is enzymatically catalyzed and one of the byproducts of this step is pyrophosphate; how can I eliminate the pyrophosphate background, because up until now I have been encountering a giant phosphate wall and the absorbance readings have been way off scale?
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If there is a buffer that has been commonly used by other researchers for this enzyme, you can start with that.
If not, the buffer should be optimized for the particular enzyme you are using. The optimization can include choosing the identity and concentration of buffering compound, the pH, the identity and concentration of salt, the Mg2+ concentration (ATPases require Mg2+ for activity), the reducing agent identity and concentration if needed, the detergent identity and concentration if needed, and a stabilizing excipient identity and concentration if needed.
Prepare concentrated stock solutions of all the ingredients you plan to test, and prepare buffer solutions by mixing them together and diluting with Milli-Q water to get the desired final concentrations. This gives you the flexibility to try many different combinations.
You also need to consider the substrates (one of which is ATP), and their concentrations.
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Working on purifying a DNA binding enzyme, pI in the range of 9. Losing it in flow-through on cation exchange, considering another type of affinity column, either Cibacron Blue or Heparin column to eliminate bound DNA.
Wondering if anyone has experience with or can comment on relative efficiency of these two media for purifying DNA binding proteins? Greater success with one vs the other? Comments and suggestions appreciated.
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I have used a library of 96 cibacron blue based affinity dyes. I guess your target DNA may bind with one of the dye. These psedo affinity dyes and you need to screen first. Let me know if you need the full text or if you have any other query.
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What factors affect the binding of endotoxin(LPS) to protein?
How to solve the problem of protein precipitation in the process of removing the endotoxin?
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Hello Xinxin,
Endotoxin can be removed with three possible mechanisms:
1. Affinity based : Affi-Prep Polymyxin resin from Bio-rad
2. Charge based : MacroPrep High Q ( When your Protein have basic pI and operating at neutral pH) , You can try Ceramic Hydroxy Apatite also.
Selecting right resin for endotoxin removal :
3. Size based : If your desired protein size less then 5kda then you can try TFF (10kda) in flow-through (permeate mode), Endotoxin have higher size so endotoxin will retain in retenate and your product will come in flow-through.
Please contact me for any more info or support.
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I have heard some rumors that the HRV-3C protease can recognize and cleave ENLYFQG peptide, which is a consensus sequence cleaved by the TEV protease. Can anyone confirm or suggest some literature where I can find this information?
Thanks in advance
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LEVLFQ\G-P HRV 3C
ENLYFQ\S TEV
HRV 3C has a strong requirement for Pro. I guess it may work if you added Pro to the TEV sequence.
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Dear colleagues,
I have a therapeutic peptide, that degrades quickly in mouse serum stability assay but has a relatively high circulation time in vivo (mouse). The cleavage site I found is suspected to be specific for thrombin, a protease that is activated during clotting and therefore active in serum but of course less active in vivo. I would like to confirm this assumption by testing the peptide stability towards pure thrombin.
Now in my lab I only have human thrombin, but no mouse thrombin available. I am not sure if it would still make sense to do the test with human thrombin, although I did all my previous tests with mouse fluids.
The crucial question is: How does the activity of human thrombin differ from mouse thrombin? Would the results still be comparable?
The MEROPS (protease) database lists both proteases under the same entry. Sequence homology is quite high, just few substitutions are present. I only found one publication that directly compares activities towards certain endogenous substrates [1]. Specificity was the same, just reaction rates differed towards some of the substrate. Can you recommend me any database, publication or other reference that would help me to adress my problem and justify my decision? I would be happy about any advice about this.
 
Kind Regards
Roland Böttger
[1] Bush et al. (2006). Murine thrombin lacks Na+ activation but retains high catalytic activity. J Biol Chem, 281(11), 7183-7188. DOI: 10.1074/jbc.M512082200
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In the end I just used human thrombin for my studies. It worked all fine. My peptides very cleaved in both mouse serum (containing lots of thrombin, as coagulation is activated) and pure human thrombin. Basically, I used the human thrombin cleavage test to verify that thrombin is the peptidase responsible for degradation of my peptide sequences in mouse serum.
The results are not quantitative though.
I think it can be justified to compare human and mouse thrombin, as the substrate specificity is highly conserved among mammals anyway. You can look this up in the MEROPS database. Also, most studies in life science aim for application in human anyway, so it's always good to have animal and human data.
Finally we published two papers including the human thrombin stability (in both of the supplementary materials):
[1] Böttger R, Hoffmann R, and Knappe D, Differential stability of therapeutic peptides with different proteolytic cleavage sites in blood, plasma and serum. PLoS One, 2017, 12, 6
[2] Böttger R, Knappe D, and Hoffmann R, PEGylated prodrugs of antidiabetic peptides amylin and GLP-1. J Control Release, 05/2018
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I prepared a solution of Pesin in 0,1M HCl and I stored it at -20ºC. Due to the concentration of pepsin that I need in this solution, I need to prepare a lot of solution in comparison to the amount that I need to use each time. As far as I know, HCl is activating Pepsin, do I need to prepare the solution every time I need to use it or by aliquoting it and storing it at -20ºC it is ok? Is pepsin loosing its activity when stored in HCl?
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Hi Anna,
I have “P6887 SIGMA Pepsin from porcine gastric mucosa lyophilized powder, 3,200-4,500 units/mg protein” Do you know how can I make the stock? I will be very great full if you help me.
Thank you so much
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Hello everbody
I checked the palmitoylation using ABE assay  for my intrest protein last year it was success but this year I repeat the assay more and more times but the problem iget every time thick band with -NH2OH although it must became lower than +NH2OH .
any advice to rid of this band
thank you
Nesma
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Well as suggested by Matti and Raquel, it may happen because of non specific biotinylation which in turn could happen
1. when thiol blocking is incomplete, I would recommend that you use overnight blocking with NEM assisted with 2%SDS . I would recommend you to used upto 50 mM NEM.
2. Concentration of HPDP biotin is too high
Preparing fresh Hydroxylamine is very important also.
I would set up a control reaction to see if protein of interest has any inherent affinity to bind to streptavidin agarose.
I would also compare silver stain profile of -NH2OH samples with +NH2OH samples to see if the effect is global or limited to given protein of interest.
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Hi,
I'm a beginner to modeller and I having trouble running the DOPE score script. It says "No module named modeller" in IDLE.
Thank You,
Mikhail
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Try to open modeller as : Run as administrator
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Why isn't serine's side chain ionizable but cysteine's is? Is that related to the different nucleophile strength between Sulfur and Oxygen ?
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Thank you both so much !
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Dear friend,
I am wondering any friend has tried/is trying to use BioID methodology to screen for proximate proteins by use yeast. If so, or if you know something about yeast BioID, please may you share the knowledge with me?
Thank you very much.
ziguo
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One group recently had suggest with BioID in yeast:
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Currently, we found one protein forms the dimer on the cell surface, but it is still unclear whether it is homo-dimer or hetero-dimer. Therefore, I am planning to use the cleavable cross-linker to cross the protein of interest. However, it seems that there is no protocol to reverse the cross-linked proteins.
Could anyone have experience on this technique give me some advice for this matter.
Thank you so much for all your help.
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Heating your SDS-sample at high temperature (95 C) for 5-10min.
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For zymography
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thank you Alim Seit-Nebi and Steingrimur
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Dear colleagues,
I detected some proteins in a tissue but the gene mostly expressed in another tissue. I guess the protein might be transported from the original tissue. How can I prove it or how can I track the protein movement in plants?
Have a good Chinese new year.
Zha
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Thanks Tomas, I am indeed looking for a protein labeling method and also think about other possibilities.
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I want to calculate the binding affinity of a protein (at a specific site) to its ligand. It is already known which atoms of a protein-ligand complex form bonds. So how can I calculate the binding affinity in this case ?
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Hi Mubasher, may I suggest you taking a look at the web application Kdeep, available free of charge at the PlayMolecule suite (www.playmolecule.org). Convolutional neural networks are used to predict binding affinity (Kd and binding free energy) for a set of docked protein-ligand complexes. Therefore, since you seem to have the binding pose of the ligand, you can input the ligand in its docked pose in SDF format, plus the protein in PDB format and Kdeep will return you a predicted affinity. I hope it is useful for you :)
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Using shRNA I knocked down a nuclear encoded mitochondrial protein from mitochondrial complex II (SDHC). Unfortunately I see a decrease in RNA and Protein expression in the Emty vector control compared to the parental cell line (no shRNA treatment). 
But when I measured the respiration of my parental cell, the shEmpty and the shSDHC cells, there was no difference between the parentals and the shEmpty vector. Indeed I saw a significant reduction in my shSDHC treated cells.
So I see a decrease in RNA and Protein levels in the shEmpty cells but apparently the protein is still functional (Same respiration as parental cells).
If anyone can come up with a suggestion or explanation I would be very happy.
Thank,
Chris
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Hi Brenda,
It is interesting that the same vector you are using is now causing loss of proteins in your cells. What I am understanding from your situation is that GFP/MOCK transfections do not cause that problem in your case. This is an indication that the vector you are using to carry the shRNA may (or may not) be part of the problem. One possibility would be the cells you are using, did you get new batches from a different source? It could also be the passage number so perhaps the longer/earlier passes may work best.
Just to be safe, make sure there is no contamination between the two prep tubes that may have occurred recently (target shRNA vs. control TRC shRNA) by sequencing, or PCR amplifying your target shRNA sequence and see if you get bands in your TRC control.
That being said, I’ve always just compared my knockdown to control transduction which was an shRNA against Luciferase protein. This is because a transfection with a certain plasmid (or a virus transduction) can alter the expression of certain proteins and the best control would be a similar plasmid but with a control sequence as you mentioned. If there is a slight knockdown with the control then you would just do relative expression. I don’t think this would hurt your experiment unless the control knockdown is substantial as in your current situation. In that case you just change the vector and/or the control sequence.
I am interested to see how the viral transductions will work out. In my opinion stable cell lines allow you to see the real efficiency of your shRNA compared to transient transfections on which maximal protein knockdown may happen 1 or 5 days later depending on the target half-life. You also get a ton more cells to work with in a stable cell line and more reproducible experiments.
Regarding DNA stability in Ecoli (i.e. avoiding recombination). I’ve never had that problem with the lenti/retroviral vectors I’ve worked with. The vectors were carried in XL-10 Gold cells that are easier to transform and they are RecA negative which reduces the changes of recombination in Ecoli. Also, I’ve always prepared plasmids as well as recombinant protein by inoculating broth directly from a frozen glycerol stock (i.e. no thaw). In other words, I don’t re-plate and pick out single colonies since technically the frozen stock was a single colony. I think in certain situations with low protein expression or a very low copy number, inoculating from a re-plated colony might work but it is not always necessary. I never had any issues with this method regarding plasmid quantity, purity, or protein expression. As far as DNA, you can keep it in the -20C indefinitely.
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I have used a protocol from C. L. German and C. L. Howe (2009) with some modifications. To be brief, I didn't use an ultracentrifuge at the end with the cytosolic fraction. I stained for histone H3, HSP70 and for b-actin. I see H3 only in nuclear fraction, but HSP70 and b-actin in both cytosolic and nuclear. My cytosolic fraction can have some of the small vesicles fraction proteins, but that is not a problem in my experiment. I need a good marker that is only in cytoplasm and not in nucleus/ER. I know that my cytosolic fraction is free from nucleus contamination, as there is no H3 there, so I feel okay about that protocol, but I can't imagine a situation in which I would have almost the same amount of cytoplasmic proteins in my cytosolic and nuclear fraction. I was thinking that b-actin will work great and now I don't know if it can show up in nucleus. 
I will be grateful for any help and I hope I made myself understandable. 
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Sorry for late reply.
I used GAPDH, Alpha tubulin, and Beta actin as cytosolic markers and Lamin A/C, and Histone H3 as nuclear markers for my cytosolic and nuclear fractions. I got intense bands for GAPDH, alpha tubulin and beta actin in my cytosolic extract. However, i saw faint bands in my nuclear extract too with those cytosolic markers. That might be because of the presence of little bit of cytosolic fraction in my nuclear extract. I was mostly satisfied with alpha tubulin and would recommend that. On the other hand, I obtained very nice, intense bands for Histone H3 and Lamin A/C for nuclear fraction without any bands in cytoplasmic fraction. I was mostly satisfied with Lamin A/C and recommend that.
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I am looking for methods that can help me in identifying natural small molecules binding with my protein of interest.
Thanks in advance.
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Hi there,
if you have absolutely no idea which binding partner you are looking for, the major two issues will be the limited amount of human material and the diversity of potential binding partners (amino acids, peptides, carbo hydrates, metabolites etc.). Therefore, you probably need a method that combines separation, analysis of binding and substance identification.
Some groups have really cool equipment for this, I think this combination of surface plasmon resonance (SPR), HPLC and MS/MS ist awesome:
...but there will not be so many labs that can do this kind of analysis.
Best,
Sebastian
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Hello
I have done a cloning and after that i have stored my recombinant E.coli cells in the -80C and -20C as a cell banks. after i have used the cells for expression i saw a basal expression in the non-induced culture(T0). after a long time storage of the cell banks in the freezer i saw that the basal expression increased. I don't understand why this ?is happening
I even digested the plasmid from the gene and cultured them again but i saw basal expression.
Another thing is when i cloned the gene into the plasmid pET32a(+) and transfered it to E.coli Rosetta-gami in the cloning steps. after Colony-PCR i saw that all of my colonies have the gene but when i induced them some colonies expressed protein in different size and some of them expressed the protein at the proper size. it is nonsense.
Could you please help me with this?
Any publications?
Thanks for your valuable suggestions.
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Hi Nii,
You didn't say what strain of E.coli you were using-if it's not recombinant deficient then you should not make glycerols in them as they can modify your plasmid over time depending on the plasmid and strain. This might give you protein the first time and either nothing or truncated proteins after that.
Store your DNA in a recombinant deficient strain and miniprep from those. Then for expression I prefer to do a fresh transformation into 'B' strains every time and use colonies from the plates within a week.
If you are using a lac promoter or the T7 system you can also block basal expression by the addition 1% glucose to your plates and media (unless you use AI media as Lisandra suggests-then add glucose only in your 'starter' culture and omit it when diluting your cells into the AI media). In the T7 system the LysS strains (again as Lisandra suggests) will also inhibit 'leaky' expression by inactivating the T7 polymerase.
Good luck
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I am doing it upto 6 hrs kinetics at an regular interval of 30 mins..it is reported that it used to attain plateu around 180 mins..checked all parametres.i am doing it on 7.4 pH scale and in dissolving it in tris hcl buffer..keeping it at 65 degrees, any suggestions?
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Hi Atul,
I'm also trying to make BSA fibrils, and having the same problem you had, could you ever resolve your problem, or do you have any updates on BSA fibrillation?
Kind Regards,
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How can I estimate Kd value or any parameter showing the affinity of protein-protein interactions, If I have measurements(band intensities from SDS PAGE) only in linear range? Any methods, other than using saturation binding curves with Prism GraphPad?
          
 
 
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You really should demonstrate saturation. Otherwise it's hard to be certain that you're observing specific binding. Maybe you can use a lower concentration of the capture/receptor protein to facilitate a higher ratio.
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Dear All,
I have read many questions and answers in this website, and found out that some people mentioned a method to reprobe another primary antibody of different target protein without stripping, but didn't mention the detail. This is very helpful, since I have bad expierience with stripping, and if two target proteins' molecular weight are too closed to cut a membrane.
Could you kindly give me a detail how to do it? for example, Do I need to block the membrane again or just incubate with another primary antibody? Must the primary antibodies be different species for this method?
Thank you
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Hi,
If you are using HRP antibodies, you can bleach the membrane with 30% H2O2 for 20 minutes. We do this for our streptavidin blots since the association is so strong between biotin and streptavidin, but this will work for any HRP-conjugated antibody to completely remove the signal. However, the primary antibodies should still be different species. You won't need to block again after bleaching, although you will definitely want to do a couple washes to remove residual H2O2. If you want to be extra thorough, you can put ECL on the blot and image real quick to make sure the blot is fully bleached, but I've never had a problem. And even if your first antibody binds to a specific protein, after bleaching you can probe with another antibody against a different epitope of the same protein and you should be able to see signal.
Hope this helps!
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I am running a 10 ns NVT production run MD simulation of a protein in NAMD. Everything looked fine till 9 ns, but during the end of the simulation, while running the dcd in VMD, it seems like the protein is trying to come out of the water box. I don't know what's the issue in here since I think I have set everything fine in the conf file. Will this effect my simulation ? and how can I fix this issue of my peptide coming out of the box ?
Thanks
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Does shifting the centre of the water box with "-shiftcenter {x y z}" do anything?
For example, a command like:
pbc wrap -compound fragment -center com -centersel "protein" -all -shiftcenter {0 1 0}
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I have a series of histidine-containing small peptides and I would like to measure their affinity for binding nickel. Does anyone know of assays that measure nickel binding? I've been able to find a ton of protocols for iron binding, but nickel protocols seem to be scarce.
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Dear Erica and Krassimira, may you send me these articles as well? Cheers.
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Hello,
I would like to work with the second virial coefficient (A2 or B22) to determine the best conditions to formulate protein buffers and also to determine the best conditions for crystallisation tests.
If I well understood, a range between -1.10E-4 and -8.E-4 increases our chance to get crystals.
However, is there a range or a rule of thumb for the aggregation value with A2 and for the best conditions for formulation. 
Best,
Sébastien
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You might find it easier to work with KD (diffusion interaction parameter). In comparison to B22, it is relatively easier to determine KD using DLS or other experiments that measure diffusion in protein solutions at different concentrations. There is more data available in literature on KD than B22, especially for the antibody solutions. 
I hope this helps.
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I have tried to dimerize this protein without success, both proteins are purified
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Thanks for the information, I will try some of them, I already have my purified proteins so I do not need to dialyze
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i have purified protein of 4.3 mg/ml. i want to study the unfolding of protein with increaing con. of urea and azide salts.  Can any one please guide me how to select protein con. to study?
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Are you looking at tryptophane fluorescence (wavelength shift or quenching, e.g. in antibodies? Whatever method you are using, your protein concentrations need to be such that you are well within the sensitivity range of the method. Make a first estimate based on the molar concentration of your protein and the number of Trp residues in your protein, then measure the spectrum for the fully native and the fully unfolded state. e.g. for observing the unfolding of antibody domains, single chain and Fab fragments, we used final concentrations of 0.2-0.5 micromolar in equilibrium denaturation experiments,
see:
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hello;
I have expressed a recombinant protein in E.coli as inclusion body. I've purified my protein by pH gradient on Ni-NTA resins and now I need to do a stepwise dialysis to refold my protein. Following this step, I'me going to inject it as an antigen to mice to generate antibodies.
Does anyone have any practical protocol or experience about it? 
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I suppose that your inclusion bodies have been solubilized by urea or GuHCl.
There is no universal protocol. This will depend on the characteristics of your protein, if it’s a transmembrane protein, a protein which is secreted (which may have disulfide briges), a protein which remains in the cytosol (which is a reducing environment), a protein with a single domain or a protein with several domains.
It is worth knowing that some proteins are extremely difficult to refold from inclusion bodies. 
There are however some common principles. First, you want to get rid of the denaturing agent. In general, this is performed by dialysis against an appropriate buffer. Sometimes, it is sufficient to lower the concentration of the denaturing agent by simply diluting your protein solution into fresh buffer. Sometimes, the dialysis has to be rather slow.
The nature of the appropriate buffer can be determined from what you already know from your protein and its natural environment.
A reducing agent (such as DTT at a concentration in the mM range) is probably required to avoid the formation of undesired disulfide bridge.
Does it contain a divalent ion such as Ca++?  If yes, the ion needs to be present in your renaturing buffer.
Does it require a cofactor which may help correct folding?
The pH may be also an important factor. Try at pH 7.5 but you may go up to pH 8.5 and down to pH 6 or even less (if your protein is naturally in an acidic environment).
The temperature may play a role but, first, try at 4°C.
Finally, you may try the refolding  at several protein concentrations. By working at lower concentration (around 50 µg/ml), you may avoid protein aggregation.   
At the end, it would be good to check the correct refolding of your protein. If you have a biological activity (enzymatic activity or ligand recognition) and a related assay, this is the best way to check the correct refolding of your protein.
If such an assay is not available, check the turbidity of your sample, spin out aggregates and see if you still have a good proportion of soluble proteins. Then, put this “soluble” fraction on a gel filtration column in order to verify if the elution volume is compatible with the expected molecular weight of your protein.   
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hi,
i want to biotinylate a fusion protein which consists of two parts. For me its important to know which part of the protein got biotinylated. How can i do that?
i only thought about to cleave the protein an do a western blot with anti-biotin-ab
but the protein has no cleavage site between the two domains.
thanks for your help
anne
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If you are working with a purified protein, you could digest it to completion with trypsin and compare the LC-MS peptide maps with and without biotinylation, then determine the sequence of the identified biotinylated peptide(s) by MS sequencing. You will need the help of mass spectrometry experts for this work.
You could also use strepatividin beads to pull down the biotinylated peptide(s) and sequence the peptides that bound to the beads.
You could experiment with different proteases, such as chymotrypsin, AspN and GluC, to try to find one that cuts with a useful degree of specificity between the two domains, based on SDS-PAGE. Also, cyanogen bromide can be used to cut selectively at Met residues.
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I have checked the peroxidase activity of E.coli under the stress of silver nitrate (20 μg/ ml). Surprisingly, I am getting negative results means control sample (E.coli without any silver nitrate treatment) found to have higher peroxidase activity silver then treated bacteria. Can anyone suggest what could be the probable reason behind this kind of result ?
Please provide me the reference if you can.
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I have used lower concentratons also varied from 1.5 microgram to 20 microgram/ml and all the reading are less then control.
plenty of reports are there, which suggests that silver used to exceed the proxidase level.
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I am doing the thermal shift assay to study protein unfolding with quanstudio 7 flex.
The following is the conditons i have tried.
1) SYPRO ORANGE, i have tried 50 X, 500 X, 1000 X, 2000 X.
2) protein concentration from 0.1 mg/ml to 5 mg/ml, and my protein is 110 kDa.
    Always centrifuge the protein sample at 18,000 g. Buffer for purifing the protein is 20 mM Tris-HCl (pH 8.0), 300 mM NaCl and 10% glycerol.
3) the buffer is 20 mM Tris-HCl (pH 8.0), 300 mM NaCl.
4) Excitation and Emission is set  to 470 and 586 nm.
5) the sample volume is 20 ul.
The problem is that initial fluorence signal is always too high, even higher than the  signal at 90 oC. The figures i attached are the abnormal melt curve plots.
Now i can not find the cause of the high initial signal. I am so confused.
Thank you for you help.
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I agree with Jaime. Your final concentration of Sypro orange is too high. We normally just use 0.5x - 1x final concentration and this gives us a low baseline signal.
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After precipitation with ammonium sulphate at 40,50,60,70 Percent saturation, I dissolved the precipitate in phosphate buffer and performed enzyme assay but did not get any activity. I want to know whether I have dialyze to the enzyme before assay or not? Cany any one suggest me, what happened wrong.
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Ammonium sulphate precipitation needs to be done on ice (not in the cold room, which - as Fleischer wrote in his book on protein purification - chill the operator, not the sample), with constant stirring and slow addition of AS (literally crystal by crystal) to avoid local high concentration. Even then, some proteins do not survive this procedure. Proteins are like people: every fool is different.
After AS precipitation the excess salt needs to be removed from the redissolved protein to recover full activity (e.g., by dialysis or desalting). Salts lower the effective water concentration and enzymes need water as "grease" for conformational changes during their cycle. Indeed, this is how AS precipitation works in the first place.
Acetone (also ethanol and methanol) can be used to precipitate some enzymes in the active form, but you need to be even more careful than with AS to avoid irreversible denaturation. Because mixing water and org. solvents generates heat, the solvent needs to be cooled to -10 °C and added VERY slowly. After centrifugation of the precipitate at -5 °C the pellet is lyophilised to remove the solvent. The resulting "acetone-powder" can be very stable if stored dry and cold (-80 °C). Again, success depends on your enzyme and is not really predictable.
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What are the differences between the structures of human hemoglobin and bovine hemoglobin?
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I hope you can find answer in these papers
Conference on Hemoglobin, 2-3 May 1957
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I am doing purification of a mammalian leucine-rich repeat (LRR) domain fused with MBP and His6 tags. But at the end of two affinity chromatography steps (IMAC then Amylose), and checking with Western Blotting (against His-tag), I saw multiple bands. So the protein is degraded.
The only inhibitor I put in my lysis buffer is PMSF.
Can anyone give me advice regarding this problem? And if anyone knows of a database or resource where I can look at the endogenous proteases found in Rosetta, it will be appreciated too.
Addition: I am suspecting that the recombinant protein is now properly folded in E. coli so I am thinking of trying expression with insect cells.
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You could try using a protease inhibitor cocktail (EDTA-free for IMAC) instead of just PMSF. Run through the purification as quickly as possible, and make sure to keep everything cold throughout.
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Am getting a chaperone at around 63 kD . I have my construct in pETM41. Am purifying with amylose resin and getting a chaperone after affinity. Will HIC after first step clean my protein?
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It sounds like you are purifying a recombinant protein in a cell-line that has chaperones to help with folding. If your fusion protein and chaperone are tightly associated, no purification technique will work unless you dissociate the complex (they will co-purify). Try adding ATP (5mM) to your column wash buffer to release the chaperonin:fusion protein complex before eluting your fusion protein from the amylose resin. In the past, I have encountered some recombinant proteins that are not released from chaperones even in the presence of ATP and needed to add denatured E coli lysate plus ATP to break apart the complex.
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i regulaarly grow in 28 degers in wild type but i mutate alanine to cystine if it grow in 28 it goes to pellet 0.5mM IPTG 5h after that i cheaked 18 degres 0.1mM IPTG 12h the yield little bit increases but not that much how to exprise more in supernatent
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If I get it right, you successfully expressed the wildtype protein at 28°C and purified it. Then, you generated a mutant construct, which is not soluble, when you express it at 28°C and only a bit better when you express it at 18°C.
This simply tells you that your protein is not that happy about the mutation. ;)
I would try to reduce the temperature even more (16°C) and express the protein for a short time (e.g. 6-8 hours). It might also help to use a different growth medium, for example TB instead of LB. You didn't write about your lysis buffer, but maybe changing this could help. Also, you could also a different tag (such as MBP or GST) or a different E. coli strain (e.g. SoluBL21, Origami).
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I have a cysteine-rich domain in my recombinant protein. results of SDS-page in reducing and non-reducing condition improved the presence of oligomers caused by disulfide bond between monomer protein in sample. the tryptophan emission spectra of my protein shows two peek in 330 and 380nm. is that true to attribute second peek to the oligomers? 
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I think this is more bigger to consider a shoulder. but I'm not sure. i  will send you .Cary Eclipse fluorimeter.
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Greetings all.
We are looking into following the kinetics of short dsRNA melting and about using a dye that will bind differently to dsRNA and ssRNA. Any idea for such a dye? Or to another simple approach to such measurements?
Yoram
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Just about all the nucleic acid-dependent fluorescent dyes bind more tightly and have a greater fluorescence increase when binding to double-stranded nucleic acids than single-stranded. You could use PicoGreen or RiboGreen or SYBR Green II, for example.
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I was inducing with 0.8 mM IPTG and kept 16-18 hrs post induction at 25 degree..am trying to reduce the temp and check the expression ; also lowering the salt and increasing imidazole concentration in the lysis buffer. Please let me know any other possible ways. How much ATP could I add for washing to break the interaction?
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You can check the attached paper.  You have to optimize ATP and MgCl2 concentration.
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can acidic pH and guanidinium hydrochloride and high temprature cleavage the protein intermolecular disulfide bond ? 
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Not cleave disulfide bonds by heating  or in acidic pH
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three truncated forms of Chondroitinase ABC1 were designed and expressed to evaluate which enzyme variants have higher/lower stability. some amino acid deleted including tryptophan and other amino acid that play a role for conformation of enzyme which could be detected with intrinsic, quenching fluorescence, limited proteolysis and Far-UV CD.
we want to explain structure differences between enzymes by intrinsic, quenching fluorescence, limited proteolysis and Far-UV CD data. considering different numbers and remained type of amino acids in each variant how it would be discussed? are there any similar articles or other methods to compare structure of those type of different enzymes? 
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You can model your proteins (WT and truncated proteins) using servers like ITASSR, HHPRED (MODELLER) etc. and then you may get some clues. Zymography is an assay in solid phase but spectroscopic assay is in liquid phase (could be different mobility of catalytic loops!!) 
I will be more then happy to help.
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I have GST  tagged  purified recombinant proteins and would like to go for CD analysis. Does someone have the sample preparation protocol for running CD analysis? 
Also, can I use GST tagged protein for CD?
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Hello Heide,
Thanks.
No I don't have ant preference for GST tagged protein. I just wanted to gain the information. Yes, now I have got the idea about cleavable GST.
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I am wanting to use a non-reducing sample buffer to resolve ubiquitin~E2 ligase interactions under various conditions by SDS-PAGE.
Looking at the literature some groups use SDS sample buffer to prepare their samples, whilst others use LDS sample buffer.
Why is this the case? Is there any significance to the cation or is it just user preference?
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Hi Hamish,
Even though there is no major difference between them, both are anionic detergents, LDS (Lithium dodecyl sulfate) is a better detergent in comparison to SDS (sodium dodecyl sulfate) if the protein is to be resolved at low temperature. As Chris mentioned, LDS buffer is preferentially used with the Invitrogen gel systems. Based on your availability you can use either!
Best wishes,
Pramod
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I wanna know how much homodimeric form or heterodimeric form there are, and I want to separe their.
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Gel filtration may not be the right approach if the masses of the homodimer and heterodimer are similar. Ion exchange chromatography or isoelectric focusing may be better approaches.
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Hey guys,
I'm having some difficulty purifying my 46 kD GST fusion protein, specifically, I'm having issues with proteolysis and purification. I appreciate all suggestions, however time and money are not on my side, so the simpler the fix or idea the better.
Induction:
To begin, I pick a colony from my colony purified GST fusion plate (transformed into BL21 cells about 3 months ago) and place it into a 20 mL O/N starter culture. Also, I'm wondering if the age of the E coli could have anything to do with proteolysis. The next day I add the starter culture to 180 mL of LB+ AMP and induce at OD600~.2 with 1mM IPTG, 37C for 5 hours. After induction I spin down my culture at 6000 x g 4C for 30 minutes. 
lysis:
On day 3 I thaw my cell pellet on ice for about 15 minutes and resuspend my it in my lysis buffer 5mL/ cell pellet(50 mM tris pH 7.5, 100 mM NaCl, 5% glycerol, 1 mM DTT, .1% triton X-100, .5M EDTA and 1 pierce protease inhibitor tablet per 10 mL lysis buffer all in PBS). I also add 1 mg/mL lysozyme and incubate 37C for 1 hr in a shaking incubator. I then sonicate my lysate on ice using a branson sonifier 250 (Output control: 6, Duty cycle %: 60%) for 15 pulses then off for 1 min ten times. Afterwards the sample becomes very viscous. I add DNAse 1 10U/mL and 1mM MgCl2 plus an additional 5 uL of 1M MgCl2 / mL cell lysate and let that sit on my bench at RT for 30 minutes with occasional shaking. After I sp