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Kindly Provide references also
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I do have a similar question Though I tried Hitchcock equation with what seems to be a better result, I still had -ve binding constant. Have been able to solve it? Harpreet Singh
Thanks
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The protein is an enzyme cloned in pET28a vector. I have used IPTG conc.(0.5mM, 0.8mM, 1mM) and culture at different time point (4h, 6h and overnight). I checked its induction 6 months back when the protein was cloned and I saw the induction.
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If you are getting the desired band in an uninduced sample also, then that might be due to leaky expression. As many articles suggest to add 1% glucose. I tried but didn't work in my case, if you want, give it a shot.
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Suppose i got two vials in which one sample is a protein (Which is a di peptide say for example Insulin) and another sample which is an Antibody, Here I've forgotten to label the samples so they got mixed up, So how to identify which sample is protein and which one is an antibody ?
Initially I thought of running SDS-gel, Both reducing and Non-reducing, But i may end up getting 2 bands in reducing condition for both Antibody (due to Heavy and light chain) and Protein sample (due to two peptide chains) and One band I'll get in non reducing condition for both cases, So this method won't work, know what to do ?
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The molecular weights of the bands, either on a reducing gel or a non-reducing gel, will be enough information to distinguish the two proteins. On a reducing gel, immunoglobulin G will have bands of 25 and 50 kDa (150 kDa on a non-reducing gel). Insulin will have 2 very small bands that will run near or at the dye front on a reducing gel, depending on the type of gel. The total mass is only 5.8 kDa.
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I know the approximate folding and unfolding rate constants, but not the exact concentrations or anything like that. Is there a way to determine a very coarse approximation for the relaxation time when the temperature is increased 2 degrees celcius? Two minutes or ten minutes or an hour?
Kfold=10^5.8
Kunfold=10^-4.1
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About the Deborah Number ― Referring to the Bible, Prophetess Deborah sang: "The mountains flowed before the Lord", celebrating victory over the Philistines. It is now scientifically established that mountains indeed "flow", considering the geological time scale. There are two different perspectives of time to be generally considered when a transient relaxation phenomena is observed; one that is intrinsic to the phenomena, another that relates to the experimental time scale. Born as a rheological concept, the dimensionless ratio between the 'time of relaxation' and the 'time of observation' was named Deborah number (De). References: M. Reiner, The Deborah Number, Physics Today, January 1964, p. 62; https://physicstoday.scitation.org/doi/10.1063/1.3051374?journalCode=pto The following reference discusses the concept of Deborah number with regards to the glass transition:
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I am purifying a recombinant protein in E.coli, with an expected molecular weight of 17 kDa and a PI of 4.6. When I put my protein sample on gel, I see a band appearing around 30kDa, so quite big but this is not the mayor issue here (although input is always welcome): This 30 kDa band is visible when I run my SDS-page with Mini-PROTEAN® Tris/Tricine gel (4-20%). When i run the same sample on a Criterion XT gel with Bis-Tris-HcL MOPS buffer system, i see this band arround 24 kDa. Any idea why this could be? I also have another protein with similar PI (and expected size of 40kDa) that shows a band around 50kDa in the Tris-Glycine but around 40 in the Bis-Tris MOPS.
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Nah, termini are almost always disordered, the rest seems fine. Also, I believe the charge would explain only why it doesn't run as it supposed to (17 kDa), not why it runs differently on 2 different gels. Other reasons why it doesn't run at 17 could be some PTM, type of buffer used, whether protein is in detergent, generally shape of a protein in case it doesn't get totally denatured and so on. Anyway, I've seen different markers giving different masses before so I wouldn't be surprised if that was the case. Unless your lab used both of these markers for a long time and they were always fine.
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Hey,
I am thinking of extracting overall protein from formaldehyde-fixed cells via the "boiling" method, as described by Sadick et al. 2016, 2017 (see below), and further process the samples in the Pierce BCA assay. Does anyone have practical experiences with this? Are there any pitfalls? Especially, I am wondering about the durability of multi-well plates at a temperature of 100C (polystyrene melting point approx 240C).
Best
Sebastian
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Fixation with formaldehyde will affect your BCA quantification and you will end up getting skewed values that will affect your SDS-PAGE outcomes.
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I want to read C-terminal and N-terminal by ion mobility, but i don’t have protocol
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Negar Saeedi are you sure that Ion Mobility is the technique that you are looking for?
regards,
GB
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I would like draw protein 3-D structure? Can anyone suggest me the best software that I can buy?
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I think for drawing of protein 3-D structure, softwares such as SPDBV and PyMOL are proper.
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Hi, I am trying to draw a secuence of mussle foot proteins like the attached photo; but I would like to know what software should I use to do so? I am using Chemdraw but I could not get the curve! Thank you 
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First try to draw in chemdraw and edit it in ppt
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I would like to ask the above question in context of intact protein characterization. Is it possible to detect DTT in positive mode using 0.1% formic acid gradient, and in which conditions?
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So, what is the issue to run in negative mode? The negative mode will give you better signal/noise ratio; therefore, you will get better sensitivity. If you need to monitor other peaks in positive mode, you can make a separate acquisition window for pos and neg mode.
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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 performed circular dichroism to estimate my protein sample's secondary structures. My sample buffer was 100 mM sodium phosphate (the CD of the buffer was measured and deducted from that of the sample). Spectra were collected from 240-190 nm and the result obtained was as follows: Alpha Helix - 16.1% Beta sheets- 44.8% Turn- 0.3% and Random coils- 38.7 %
However, the CD spectra obtained had a negative dip at 208 nm and a positive one at 192 nm. Is this normal? If not, what should I do? Thank you.
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Circular dichroism is relatively good in comparing (relative) results where for example your peptide is studied at different temperatures, with or without anionic phospholipids and so on. This allows you to say “upon increase of temperature one sees a decrease in helical content”.
See for example:
Unfortunately the deconvolution of CD spectra varies a lot and depends on the method used and some tent to over-estimate the alpha helical content. So an absolute answer based on one method is not THE answer of the analysis of your spectrum. The best you can do is comparing the results of different CD analysis since numerous programs are available (often your CD equipment includes pretty good software as well).:
Dichrocalc:
Dichroweb:
BestSel:
K2D3:
Dicroprot:
There are many more different programs to deconvolute your CD signal in order to extract secondary structure contents (CDPro, CDNN etc.).
For a nice introduction see:
PS. To get some ‘grip’ on what to expect in terms of secondary structural elements you might consider to do some secondary structure prediction, like:
SOPMA
or
JPred
Hope this answers your question somewhat. Good luck!
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I've been using microBCA for protein concentration estimation for exosome samples. I used to get very consistent results with lysed exosome samples, however this is no longer the case and I am getting very low, and even negative protein concentrations calculated using this assay now.
I have thought it may have been something wrong with the lysis buffer, I normally used a 1% NP-40 lysis buffer, and have tried using RIPA buffer as well as a 0.1% SDS buffer. I've checked for incompatibilities with the reagents and have ruled these out.
I have also used a Bradford assay on the same samples, and instead of low and negative values, get uniformly high values, but when these samples are run on a gel, I see nothing appear using Ponceau-S or Coomassie staining suggesting there's nothing there.
I'm at a loss and any help or advice you could offer would be appreciated!
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Micro BCA is more sensitive than standard BCA, but the catch is that micro BCA is inhibited by the most common buffers such as RIPA or TBS, and many more chemicals. It is recommended a 1:10 dilution in ultrapure water before doing measurements with micro BCA. RIPA or TBS do not inhibit standard BCA at all, so I would go for standard BCA unless I do not have much exosome sample, which happens often.
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I am facing a problem trying to measure the secondary structure of a protein by FTIR. The protein remains adsorbed to the surface of the ATR diamond.
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Short peptides, isolated proteins and soft tissues. Using a piece of laboratory towel.
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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'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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soluble protein characterization
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Hello Shobana, i do not see where you have the difficulty because the classical methods of protein separation and characterization have not changed, at least not to my knowledge. Protein characterization in recent times involves five protocols; protein quantification (involves Absorption at 280 nm, Lowry assay, Bradford assay and Mass Spectrometry - a technique by which you can determine the mass of a protein with remarkable precision. It can detect any important modification (post translational modification) or variation in a protein structure), Electrophoresis (migration of a protein in a gel as a function of size, shape and charge), dynamic light scattering (a technique used to determine the size distribution profile of small particles in solution) and circular dichroism (a variant of absorption spectroscopy which measures the difference in absorption of left and right polarized light (in the ultraviolet) by a medium. For protocols visit the links below: https://www.uq.edu.au/pef/content/characterisation-techniques-0.
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Hello all, 
I was wondering what HSQC pulse sequences help to reduce T1 noise. We have a Bruker TCI 600 cryoprobe and have tried several different pulses, but have a very large T1 noise problem when using the Shigemi tubes.
What sorts of advantages would an adiabatic pulse sequence have in this case? Specifically, we are looking into the "sensitivity improvement" feature in some of the pulse sequences, and have only found variants that are phase sensitive, is this a byproduct of the sensitivity improvement pulse? Or is it possible to eliminate the phase selective feature with these pulses?    
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Hello all,
do you have any good experience with a pulse sequence to reduce T1 noise on HSQC without any application of gradients? (Bruker systems)
Thx...
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I wanted to check translocon proteins (Sec61 alpha, beta, gamma) in endoplasmic reticulm. I got band for alpha subunit protein. However I could not band for beta sub-unit (15 kDa) and gamma sub-unit (7 kDa).
I used following conditions, Gel Running: 16% SDS Gel, 200V, 400mA, 50 Minutes
Blotting Condition: Buffer with 20% ethanol, 80 V/gel, 400 mA, 40 minutes., Nitrocellulose Membrane.
Blocking, 5% Skim Milk, 1 hour.
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I have been looking at the translocon and use 4-20% gradient gels. I use a normal blotting protocol but fix the blot in 5% acetic acid before blocking. This works well for the beta subunit.
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What is the difference between a recombinant protein and a native protein?
Why is expression of recombinant protein may be toxic with compare the same amounts of native proteins? This problem is connected with folding in unnative organism?
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For simple protein enzymes like insulin you can use bacterial expression systems to create proteins.
For any protein that requires complex folding, and post translational modifications, you would need eukaryote expression systems eg. CHO, yeast etc, that can express these complex proteins, that require chaperones, correct tRNAs, and post translational modifications that a bacterial system can't provide.
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I have isolated and identified more than 40 hypothetical proteins from E. coli by using MALDI-TOF and LC-MS/MS. Hypothetical proteins are cloned, over expressed and two proteins are characterized by binding study and crystal study.
Most of the proteins are not forming crystals. It is very hard to make deletion mutant of all hypothetical genes and make characterization of hypothetical genes. I have isolated and purified most of the hypothetical proteins by using chromatography. I would appreciate if anyone could give suggestions and advice regarding the protein identification by using any functional studies.
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I think that is very important a refined bioinformatic study of hypothetical proteins for planning a very detailed set of wet-laboratory experiments
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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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I am making a fusion protein with E. coli enzyme ilvD and the fluorescent protein mNeonGreen and I am trying to find more information regarding what the linker sequence between the proteins should look like. 
One sequence I am considering is the [Gly(4)Ser]x3 sequence, but I am interested in more options.
I should also say that I am only interested in expression level of ilvD, so it is not important to me that ilvD remain catalytically active, only that fusion protein transcription/translation is efficient and that mNeonGreen can efficiently fold/fluoresce. 
Does anyone have any experience with fluorescent fusion protein linkers that would be pertinent?
Thank you very much!
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Nicolas, grab it from here: http://www.kazusa.or.jp/codon/
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I'm using the 15% hand casting gel for a 9 KDa protein, later a 4%-20% hand casted gradient gel. This problem the sample band became a line, and cannot be recognized which is which. It happened on every sample including ladder.
So I re-prepared all the casting buffer I used and checked with the Tris-HCl pH. Everything seems correct. After several times of SDS-PAGE, I found 10% gel, 12% gel can run nicely. But when percentage goes higher, the spread problem appear. It's hardly to believe that the casting buffer is not proper. 
The running is voltage constant at 80V. The current is 25mA for one gel and dropped to 10 mA in the end. It took 2 hour for 15% gel. I tried 80V for stacking gel, and increased to 110 V for resolver gel or 80V all the way. The spread problem is there. 
I really want to know whan happened on my 15% SDS PAGE gel. REALLY! Please give me some suggestion, ideas or anything concern that come to your mind. Thank you so much!!
This is my 15% gel and 5% stacking gel recipe.
15% gel: Distilled water 2.3ml, 30% Acrylamide 5ml, 1.5M Tris pH8.8 2.5ml, 10%SDS 100ul, 10% APS 100ul, TEMED 4ul.5% stacking:Distilled water 2.7ml, 30% Acrylamide 670ul, 0.5M Tris pH6.8 500ul, 10%SDS 40ul, 10% APS 40ul, TEMED 4ul.
The APS is freshly made before casting gel. 30% Acrylamide is mdae with 29.2g Acrylamide and 800mg N’N’-bis-methylene-acrylamide.
(I showed two pictures below, one is the western blot for beta actin. There supposed five samples, but they form a horizontal line. The next one is 4%-20% gradient gel. I randomly chose a 20ug protein sample. Loading volume is 20ul. It's clear that the lane became wider and wider as it ran towards to the bottom.)
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Hi, 
I am also facing a similar problem. If the problem is not solved yet with you, the reason for lateral spread of protein leading to fusing of bands could be that the Ionic strength of sample lower than that of gel (if other obvious possibilities are obviated) (This is something I gathered from this site: http://www.bio-rad.com/en-us/applications-technologies/performing-protein-electrophoresis) .
Let me know if this works.
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I am doing western blots to determine if antibodies are binding selectively to the desired protein in a basal fish. I am using two different antibodies for the same protein in different vertical strips. They have different immunogens. One measures 110 kDa (the predicted molecular mass), the other measures 100 kDa. Uniprot states there is a 100 kDa isoform, but this hasn't been confirmed in our species. Is a 10 kDa difference in molecular mass acceptable considering they were run on different gels and possess different immunogens? What's an acceptable kDa range for the same protein on different gels? Thanks. 
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The question is whether the measurement of 109 and 100 Da made on separate strips is a statistically significant difference or is within measurement error of being the same mass. One way to address this question is to make the measurements several times, calculate averages and standard deviations, and perform a t-test to see if the difference is significant. Another approach is to combine both antibodies in one blot and see whether you get one band or two, compared with the individual antibodies. This should be done with a light loading of antigen to maximize the chance of seeing two separate bands. Also, you should optimize the separation, if there are two bands, by choosing the most appropriate acrylamide percentage.
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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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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 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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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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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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protein, protein dimerization, buffers role, salts role in proteins precipitation, proteomics, protein crystallography
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Citrate is not only a good buffer in the slightly acidic range (pH 2-6) and a chelating agent for bivalent metals, it is also a cosmotrope (right side of the Hofmeister series of anions) that stabilises protein structure.
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I don't care if the protein conformation or biological activity is lost. I just want to change Asn and Gln to Asp and Glu. I do want to avoid proteolysis.
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We have identified a novel protein with a ricinB-like (lectin-like) domain. How can we identify the probable function of this protein? Please help.
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Hi Riddhi,
The responses above are elaborating on how to indentify the interactions of this gene. To identify the function, you gotta knock-out/off this gene and do a mutant analysis.
Thanks,
Aniket
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I am working on a protein which is having the propensity to dimerize. When using sample buffer with 5% of 2-ME, I am getting both the monomeric and dimeric forms.
Any suggestions for the final concentration of 2-ME to get the protein in monomeric form?
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Hi Avni, it could be that your  protein is dimerizing in the gel because mercaptoethanol does not travel into the gel along with the proteins because it is uncharged. The same goes for DTT.
If this is the case, then treating the sample with iodoacetamide (~5x concentration over the 2-ME) should explain if this is the problem.
Iodoacetamide will bind all free -SH groups and prevent any S-S linkage.  
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After DSF guided refolding of protein, results for denatured and refolded samples are coming out like this. is it ok if the denatured sample give peak and Tm like this? as i have used mild solublization buffer (2M Urea)
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many thanks james.....your suggestions and explination is really helpfull...thanks again
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My protein is for 12kDa, doesn't have any disulfide bonds. I have 6ml of purified denatured protein with good concentration. But have limited arginine. Can I use 50mM arginine & get increased yield of properly folded protein without aggregation? Please suggest.
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Hi Deepika 
Your protein is 12 KDa with no-disulfide bonds.
Arginine avoid dis-aggregating in refolding process but concentration depends on our hydrophobic amino acid.  The 0.02M-0.3M Arginine is preferred for more native structure of protein (depend on refolding protein concentration).
Before that you have to denature the protein with correct concentration of Urea/GuHcl and BME/DTT.
If i  know more about you protein, suggest better way. 
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Can anyone suggest me, whether primary mouse Abs (94) can be renatured after use of 8 M Urea (as by using 8 M Urea it will get denatured and I need to recover the Abs). If yes, please refer me the article or share the way how to do it.
 Thank You
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Hi Waheed, it all depends on the antibody. Some monoclonals are rock solid and some monoclonals loose activity when diluted into water.
If you have a reliable ELISA assay for your antibody then you should use it to determine any loss of activity.  
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I have a peptide that exhibits the normal alpha helical CD signature (positive peak at 190 nm, negative peaks at 205 and 220 nm, flat and nearly no signal from 235nm to 260nm) when free in solution.  After I conjugated it to dermatan sulfate, I find that it's signature has a shallower positive peak at 190 nm and deeper negative peaks at 205 and 220 nm, but a new large positive peak at 235 nm. I originally thought this was indicative of some uncoiling after conjugation presumably because dermatan sulfate has a large negative charge and that perhaps the spacer was not big enough.  I added a little longer spacer and it still looked about the same as before the spacer so I am wondering if the new peak is indicative of something else?  I had trouble finding information on what causes peaks in the 225nm to 260nm.  Any advice would be appreciated.
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Aromatic side chains do not actually have an intrinsic CD signal, because they do not possess any center of asimmetry. However, they may show a CD spectrum if they are embedded in an asimmetric environment, such as that typical of the tertiary structure of the peptide backbone. Thus, it seems that your peptide has some secondary, but no tertiary structure, into which it could reorganize as a consequence of conjugation to dermatan sulfate. I mean that the entire complex peptide-dermatan sulfate acquires a tertiary structural arrangement, to which the 235 signal could be ascribed. Take a look at the near UV CD spectrum up to 300 nm, which should not be flat, but show novel absorption bands, even if they are less intense than the far UV ones.
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Hi
I have obtained crystals of Cytochrome P450 3A4 grown in 6.5 % PEG3350, 12.5 % MPD and 8.5 % glycerol.
The reservoir has 13 % PEG3350, 25% MPD, ligand of choice i either 100 mM Imidazole;MES buffer pH 6,5 or 100 mM HEPES;MOPS buffer pH 7,5.
Protein solution has 50 mM KPi, 1 mM EDTA, 2 mM DTT, 500 mM NaCl, 17 % glycerol. I mix the solutions 1:1 µl.
I'm guessing that the conditions in the drop is not quite adequate for cryoprotecting the crystals?
Any suggestions how much and what to add extra? Do I bump up MPD concentration or add more glycerol?
Is it preferable in general to add cryoprteoctant to the mother liquor or is it fine to use a reservoir solution with more cryoprtectant if needed and then soak the crystal.
Thx in advance
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Hi Daniel,
your crystallization solution might actually be pretty close to being cryoprotective. Both glycerol and MDP are cryoprotectans. You can test this by freezing a drop of the solution (without a crystal) in a loop and check if it remains clear, which suggests its cryoprotected or whether it's turns hazy. If it turns hazy, I would make a solution as closely matching your mother liquor as possible and slowly increase glycerol or MDP (usually by 5% steps at a time). Glycerol usually is croprotective around 15% but with MPD in your solution you might need less. Generally it's hard to predict what works best since it is different for different crystal systems, loop size used and procedures. A good rule of thumb is to change the condition as little and as slowly (stepwise) as possible.
Good luck.
Bernhard
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My protein is recombinant human protein and its concentration is very low.
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I can recommend a new fluorescent method very sensitive that can help you, here is the information
Good Luck
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My protein forms dimer using Cys-Cys bond. I purified it using beta-ME in buffers, but not sure whether the Cystine bridge is the sole cause of dimerisation and whether if beta-ME is interrupting the Cys-Cys bridge. I analized my sample in denaturing PAGE. 
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Hi Arnab
            I will suggest you to run SDS page in non reducing and denaturing condition i.e in the loading dye do not put any reducing agent. If you protein forms disulphide linkage, you can clearly see a band near dimer of your protein of interest.  I have done this and was able to see clearly a dimer. The control for it will be a protein sample denatured with normal SDS PAGE dye i.e reducing and denaturing dye. Hope this will help you. 
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I am a beginner of western blot and my target protein if 124kDA whereas the endogeneous controls are 41kDA and 55kDA. I did wet transfer at 100V, 80mins and successfully got the band for target protein.But both of my endogeneous controls did not show any band. I thought that it might be due to the overshoot of low MW proteins. Next day I cut the gel and did transfer at 100V for 40mins for the smaller ones but this time also there was no band.But in every cases ladder was perfectly transferred in the membrane. Can I transfer the target and endogeneous control at same duration and same voltage? Why do the low molecular weight proteins not show any band in western blot?. The blots are attached below.The first two blots represent controls.
Thanking you
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Thank you so much
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We did an IP experiment and sent the results for mass spec.  I confirmed by western blot (on the same sample) that some positive controls were present -- including the antigen.  The mass spec results did not show these positive controls present in the results.  
When we IP'd against an antibody for ProteinX, ProteinX was not present in the mass spec.   Based on previous experiments we do know that ProteinX is indeed detectable by mass spec, and we also know based on Western, that it was present in our submitted sample.  Any ideas what might have happened?
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Not enough information provided to answer your question.
Mass spec's are NOT universal detectors. They do not and can not "see" everything.
For an LC/MS system to detect a compound, it must ionize well under the conditions used. This relies on many things, including, the concentration of the sample present, the experience and skill of the operator (This has the largest effect of all!), the type of MS system used, the mode the system is operated in (pos/neg) and of course, the many settings and parameters used to detect it (all of which can result in no signal, noise, signal or just about anything else).
For now, perhaps forget about the MS aspect and ask yourself if you can detect the sample using a good quality HPLC method with UV detection? Alternatively, if enough sample is available, try flow injection with the MS set up in a wide scan (look at the TIC) to see if anything is there and then a narrow SIM mode to look for fragments or clusters of the protein at higher energies.
*Most importantly of all, get some help locally with this (not the web) as a student would not be expected to do this on their own. It takes many years to learn how to analyze samples by HPLC and/or LC/MS and your focus should be on having someone help make sure you are pursuing a correct approach to solve the problem. Let them help you insure the instrument(s) and method(s) used are applicable to your sample and problem.
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i dialysed protein prdx6 in 50mM Trishcl ph 7.4 without nacl. i run sds page but i don find the protein. what should be done to get my protein. will it be useful if i dialyse it again with correct buffer i.e 5omM Trishcl+100mM nacl. 
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I agree with Omar and Bobbi, you are loosing your protein in the dialysis procedure. If you are trying to concentrate a protein solution ~below 10 ng/ml, you will loose most of it. Dialysis bags have a huge surface area that can bind proteins.  
If you can, lyophilize the solution. If you can't do that then use centricon fiters.   
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Can someone assist me with a workable procedure for determination of molecular weight of viral coat protein through SDS-PAGE? I would also be happy to know how the molecular weight is estimated from the gel.
Thank you.
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Unless there is something unusual about your protein, you can use standard SDS-PAGE methodology. To determine molecular weight, using a single acrylamide concentration gel (not a gradient gel), and a set of commercial molecular weight markers run in the lane next to your protein. Plot a graph of Rf versus log molecular weight of the standards, and use the Rf of your protein and the standard curve to measure the molecular weight of the monomer. (Rf is the ratio of the distance from the top of the separating gel to the band divided by the distance from the top of the separating gel to the dye front.).
If you are interested in knowing the molecular weight of the oligomer, then you should not use SDS-PAGE. There is a procedure to determine the molecular weight by non-denaturing PAGE, which involves running the protein on gels with varying acrylamide concentrations. You could also use analytical ultracentrifugation, sucrose gradient ultracentrifugation with size standards, and size-exclusion chromatography coupled with multiangle light scattering, all of which require specialized equipment.
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In my case, with urea fluorescenece intensity increases with red shift and with GdmCl fluorescence intensity decreases with red shift.
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How far did you go in terms of urea and GdmCl concentrations? Are you sure that both transitions are completed? Does your protein unfold through intermediate state? For many globular proteins, the use of GdmCl allows decoupling native state <--> intermediate (molten globule) state and intermediate state <--> unfolded state transitions, whereas in urea these transitions typically overlap. Also, GdmCl-induced curves are typically noticeably steeper than urea-induced unfolding curves. To verify complete unfolding mechanism of your protein you should use more than one technique. For example, near- and far-UV CD are typically used to detect changes in tertiary and secondary structures, respectively.  
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I want to work with Intrinsically disordered protein and its complexes (also a protein). In this complex I would like to use ff99SBildn for IDP and ff14SB for its partner protein. What I did is generate two different prmtop files for both the proteins (separated partner proteins into two files and used as input for tleap). Now, how can I merge these prmtop files? Is there another way to integrate two different force fields? Please suggest a way out.
Thanks in advance.
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Hi Naga,
I am curious. How do you plan to deal with the interaction between the two proteins? In my study of association between an ordered protein and a disordered one, I used coarse-grained protein models and I did change the force parameters for each protein separately to match with the experimental measurement of the isolated proteins.
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I want to invest the potentially drug-induced hemolysis of Quinpirole with a spectrometer. Therefore I want to make a standard curve for %lysis against measured absorbance of hemoglobin at 540nm. However, I need to know if Quinpirole absorbance is in that range and I can't find it in any database ...
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I couldn't find it either, but looking at the structure of quinpirole, I would not expect it to have any absorbance in the visible range. You can see this for yourself by the fact that it is a white powder. A solution of it should be colorless. Therefore it will have no absorbance at 540 nm, which is in the visible range.
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I recently purified two GST tag fragments that are below 100 amino acids and did pull down assays. However, neither of them interacted with the other purified protein. We know that the two fragments should interact with the other protein since we've done a truncation of the fragment from the FL protein and it abolished the binding. I wonder if the fragments might form soluble aggregate and it prevents the interaction with the other protein. Does anyone have any advice on this problem?
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I agree with the previous suggestions. Three main things could be occurring, perhaps more than one of these at the same time:
1. The peptides binding site was lost by cleavage
2. The peptides are self-aggregating
3. The peptides are not correctly folded and the binding site is hidden
Since you believe there is aggregation going on, I would suggest adding a small amount of a non-ionic detergent to prevent aggregation. After aggregation you would need a stronger chaotropic agent to disrupt the aggregates. Good luck.
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I have a complex of 7 proteins that I am recombinantly expressing and purifying on nickel (1 of the proteins has a his tag). I know by coomassie staining and mass spec that all 7 proteins are present. These proteins should form a ring structure, but when I look by EM I am not seeing the rings. I want to try crosslinking the proteins to each other as soon as they come off of the nickel column in case the ring is very unstable. What is the easiest/best protocol for protein crosslinking? Formaldehyde, glutaraldehyde, DMP?
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It could be, it depends on the number of lysine residues on the surface of your proteins. I prefer to run reactions with glutaraldehyde and proteins at 4C for 2 h. At 37 C the reaction rate will be fast and since you cannot vigorously stir your protein sample you need to give glutaraldehyde some time to generate homogeneous reaction conditions. Add a reduction step using sodium borohydride. 
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I am using T2 cell line. Incubation with peptide for 18 hr. Using BB7.2 monoclonal antibody conjugated with PE (conc 0.5μl in 40μl cell)(2.5μl in 40μl- 80,000 cells). Then running FACScan. 
Even in no-peptide control(-ve), there is full signal in presence of AB or in lower conc of AB, No signal in both control and presence of peptide.
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Right, I do understand that. Thank you for your comment.
But, through flow cytometer analysis, cell population that have bound antibody between control cells and the cell pulsed with peptide of interest are almost identical.
To elucidate, the question I have is should the peptide at least stabilize the MHC with small amount that we see a difference between control and peptide pulsed cells.
 
Has anyone else done T2 stabilization with peptide pulsing? A little insight would be really helpful.
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Hello, I already tested by NMR the interaction of an antibody-amyloid beta and now I want to test the influence in amyloid beta 1-42 aggregation kinetics of the presence of an antibody in solution. Due to the availability of equipments my first option is tu use ThT assay.
My main doubt is if the ThT will interact with the antibody (mouse IgG1) in solution and give me masked results.
Of course, I will use controls for assessing that hypothetical influence, but if that instruction will be enough for making useless the assay, I will try another option.
Thank you very much!
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Hi Ben, thank you.
Anyway, I am trying to monitor aggregation using ThT and I am having outrageous results. Final concentrations and conditions:
- Abeta 1-42 (pretreated with NH4OH and resuspended in NaOH (no sanitation, no vortexing): 56 µM
- ThT: 0.2 mM (in PBS)
- PBS 10 mM till 100 µL
- Greiner black flat bottom plates
Everything at 4ºC till plating and start measures (440 nm exc and 484 nm emission) each 5 min, 15 sec shaking (6 mm amplitude) before measurement, 37ºC. Equipment (tecan infinite 200) gain setting at 140.
Fluorescence values don´t increase but even go down 5-10 min from assay start and stabilize. I repeated with 3 different Abeta aliquots and I see the same. Maybe my aliquots are a bit old... but in ELISA still worked fine.
Any suggestion? Thanks in advance!
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hi i am doing a EMSA experiment,i want to know the concentration of hepain in EMSA buffer
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Hi
Overdose heparin can compete with your protein and protein/enzyme can be titrated out. So its better to use synthetic poly (dl:dc) as suggested by Bhaskar Gouda.
Best of Luck
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The sample buffer has 500mM of imidazole.
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(1) Check your amino acid content for side-chains which absorb at that wavelength: tryptophan (W), tyrosine (Y) and cysteine (C) (or use something like http://web.expasy.org/protparam/ which will give you a predicted extinction coefficient)
(2) Make sure to remove the imidazole from your sample either via a buffer exchange column or dialysis. High concentrations of imidazole will skew any data as the overall absorbance will be too high and out of the range of detection.
(3) Test your nano-drop with a different protein, nano-drops can break down or might need a good clean as the sensor area is very small.
(4) If you have access to a spectrophotometer, try using it to detect your protein instead, as these instruments tend to be much more sensitive.
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Further, if I am using a bioengineered skin construct for experiemental purposes, what is the best way to integrate a known amount of  glycated proteins into the sample?
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Thank you very much for you help Betty,
Is it possible that I could use a decellularized section of human skin, which would contain the collagen matrix as a foundation for developing my artificial skin construct?
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Does anyone know of methods to identify binding partners (e.g. receptor proteins) for GPI-anchored proteins? The protein I'm studying has an N-terminal signal sequence, which is removed prior to membrane insertion. Part of the C-terminus is also cleaved so that the GPI anchored can be attached. I wanted to use a vector using this protein fused to a Myc tag, so that I can do an IP and then send it for mass spec. However, I'm worried that the Myc tag would also be removed prior to being inserted into the membrane.
I would appreciate any suggestions! Thanks in advance! 
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Great question, its a typical problem for proteins with N-terminal and C-terminal processing, and considering the function of a protein it is relevant to all proteins that are modified by tagging with epitopes (FLAG, HA, c-myc...) or fluorescent proteins (GFP, YFP, RFP....). For a GPI-anchored protein I would seriously consider producing a recombinant secreted derivative in a eukaryotic expression system and raising specific antibodies that you can use to immunoprecipitate the native protein. However, I understand that this takes time and often people try epitope tagging first.
1) One approach would be to add the tag between the signal peptide and the mature portion of the protein, possibly with a glycine-serine linker after the epitope tag. I would also insert the first 3 aminoacids of the native portion of protein before the epitope, in order to avoid any changes to the signal peptide processing site. Also eukaryotic signal peptides are less sensitive towards the type of amino acids immediately after the cleavage site compared to prokaryotic signal peptides, it can happen that the epitope sequence is causing problems with signal peptide processing at the ER translocation pore. If the signal peptide is not cleaved, you end up with a membrane anchored protein that will probably not fold correctly. I would also try different tags to make sure you find one that works.
2) Alternatively, you could add the epitope tag before the processing site of the C-terminal transmembrane domain (the hydrophobic portion that is cleaved before the GPI-anchor takes over the function of membrane anchoring). But again, you have to make sure you don't upset the processing site, so I would insert it a few aminoacids further upstream and again I would try different epitope tags in parallel to find one that keeps the protein working normally.
In an case it is a tricky task and you have to approach it with trial and error. You are absolutely right to be worried about the removal of the epitopes if you simply insert at the N-terminus (before the signal peptide) or the C-terminus (after the hydrophobic transembrane domain, it is not going to work, don't even think of trying that. If you detect a protein, then it has not been processed properly and it is probably retained in the ER as a malfolded protein. If it is processed correctly, the tag is removed and you can't detect it. In all likelihood, both will happen, so you may see functionality in some bio-assay, and you may detect something with your anti epitope antibodies, but both methods report on different molecules so the results will be misleading. 
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It is known that N conversion factors are used to determine protein content of a nitrogenous compound. My question is if we have an amino acid, for example arginine and its N content is 32% (therefore conversion factor 3.125) and we know 1gm arginine contains 0.32g nitrogen. Then what is the protein equivalent of 1gm arginine? Is it 0.32*3.125=1gm protein or is it the nitrogen content *6.25: 0.32*6.25=2gm protein? Thanks. 
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Crude protein or stock feeds (as mentioned by Dr.Milham) may contain aminoacids,amides or non- protein -N.So a general factor of 6.25 (assuming 16% N in the protein) is used to convert estimated N to protein.However if one knows the amino acid composition of protein and its N concentration accordingly factor will vary (as per the example for arginine given by Chandini).So one can derive and use a factor if one knows the amino acid composition and N concentration in individual aminoacids.The paper attached by Dr.Weller also supports this view.
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I am using the pBAD24 arabinose inducible system in Shigella to express a protein of interest. My goal is to maintain induction of this protein during intracellular plaque assays. These assays require my mammalian cells to be plated 2 days prior to infection, then 3 days to allow for visible plaque formation. Thus, I need my cells to be glucose free for at least 5-6 days during my assay. Since this construct is repressed in the presence of glucose, I will not get any expression of my protein in typical cell culture media, which contains high glucose. Is it possible to grow mammalian cells without glucose? 
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Hi Prakash,
The cells are L2 mouse fibroblast cells. I have also tried supplementing my glucose free media with 1mM pyruvate and the cells also die. 
Jessica
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What does CD spectra at 230nm tell you about? I am working with Cu/Zn SOD which is a dimeric protein. I can see some changes in the far-UV spectra at and around 230nm. So I would like to know how an increase or decrease in ellipticity at 230nm signify?
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I agree with  Martinelli and just want to add that you should try the top two programs for deciphering percent of alpha and beta structure (one  of which is mentioned above- Dichhroweb). Remember that the curves are a combination of the percentages of different structures and thus only after doing this analysis you can get the whole story.
Gustavo
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Hi,
Yesterday i was purifying biotinylated molecules with a neutravidin polyacrylamide resin and after incubation with cell lysate, the resin looked "aggregated" instead of a conventional resin suspension.. After extensive PBS 0,5M and PBS washes, it keeped in the same way.
Do you know why this happens?
Thanks for reading
Martin
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Hi Martin, neutravidin can bind 4 biotins and if your protein has more than 1 biotin you are seeing crosslinking between the biotinylated protein and the neutravidin resin.
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I am running Analytical Size exclusion using a Superdex 200 Increase 10/300 GL column of protein samples at ~40uM concentration. The protein is known to form hexamers and dodecamers as well as other oligomers, with a monomer being ~70kDa.
I see a rounded hump in the mutant and the overlapping of peaks in the WT.
Do you have any idea what is causing this? and why the proteins are eluting at a different volume than expected?
Many thanks,
Ruth
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Thanks for the answer and protocol Omar. It will come in very useful!
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I am using DLS measurements in order to calculate the second virial coefficient of a full-length mAbs in the presence of different buffer compositions and to do that, refractive-index increment for the protein/buffer pair (dn/dc) is required.
Have you some experience regarding that?
Particularly, I would to know if I can use the dn/dc = 0.185 g/mL that is usually suggested for globular protein also in my case, where the shape is not exactly globular. If I cannot, do I need to determine it by refractometry or there is a dn/dc that can be used for full-length antibody?
Thanks
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I think you will find this paper useful:
My takeaway from it is that for larger proteins, the value of dn/dc falls within quite a narrow range, and depends on the amino acid composition rather than the protein shape.
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We observed decreased proteolysis of a Cys-containing complex in the presence of TCEP. The question is whether  this is due to the effect of TCEP upon the protein complex or on Proteinase K itself (also has cysteines).
 Thank you!
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According to this article, proteinase K contains 5 Cys residues and 2 disulfide bonds, so it is plausible that reduction of the protein could affect its activity.
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I am trying to conjugate 5/6 Fam-NHS ester to a peptide in solution. I do see a conjugated product that has a mass of my peptide +3 units of 5/6 Fam but that mass is also exact to the 9 subunits of 5/6 fam itself?
Has anyone observed that too? I know I can do MS/MS on the peak to know if it certainly is my peptide with fam units or a multimer unit of Fam?
But just as a curiosity I would like to know if that has been ever observed that before?
inputs will be much appreciated.
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Does the peptide have at least 3 primary amino groups (N-terminus and Lysine) to account for the 3:1 stoichiometry? If so, that is probably what you made.
I have never heard of self-reaction of the reagent being a problem. The dye has no amino groups to react with NHS ester.
By the way, you may find that having 3 FAMs/peptide is too many. The peptide may become very hydrophobic, and the dye molecules may quench each other's fluorescence.
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I recently solved a crystal structure for small molecule and submitted the cif file for scanning alerts, if any. I was able to rectify all the alerts but got stuck with this one. 
PLAT213 Type_2 Test ratio adp max/min in main residue(s)
The main axes values of the ADP(S) of the main residue(s) are determined and
ordered: U1 < U2 < U3. The value of SQRT(U3/U1) main axis ADP ratio (Angstrom
Units) is tested for the main residue(s). Large values may indicate unresolved
disorder. Oblate criterium: U3 - U2 < U2 - U1. Prolate otherwise.
PLAT340 Type_3 Check Bond Precision for C-C in Light Atom Structures (Z(max) < 20)
The average s.u. for X-Y bonds is tested (named bond-precision). X-Y will
generally be C-C bonds, unless there are none. In the last case the s.u.'s of
the lowest element numbers are considered (excluding hydrogen). There are
three test ranges: one for structures with the largest element Z < 20, one
for the largest Z in the range 20 to 39 and one for structures with Z(max) 40
or higher (_340, _341 and _342 respectively).
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Try to give a bigger value for omit command
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Hello everyone,
when I studied certain protein in patients who had different stage of liver disease (HCV, liver cirrhosis, HCC) and control group, I surprised that ELISA showed difference in the levels of this protein in serum but no difference have been showed in gene expression by real time _PCR and fold change equal 1 or 0.99 in all groups
so, I want to ask is there any reason that make gene expreesion the same in different groups but protein levels difference?
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Michael is right, RT data (from biopsy material?) not always reflect the changes in protein levels, especially if you are working with serum of patients. It depends on the production rate/secretion rate of your protein of interest (is it actively secreted or a sign of cell damage like the transaminases?), the stability in serum and the excretion/degradation.
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We have analyzed MALDI-TOF spectra of our peptide hydrogels and observed 6, 8 KDa molecular wt. of each but intensity of those peaks are very low. We want to perform ms/ms spectra of those low intensity peaks. Is it possible? Can anyone provide any reference?
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Hi Snehasish! If you are certain that the peptides you see are of genuine interest (i.e. you repeated the experiment and always the same types of masses appear) than it is certainly worthwhile to try different pre-Maldi workups - The quality of MALDI-TOF spectra is strongly influenced by the purity of the sample (i.e. salts, contaminants which prevent proper co-crystallisation of the matrix etc..) and therefore pre-treatment of the samples can enhance the sensitivity significantly. Luckily for peptides many tools such as C18 ZipTips or mini SPE cardriges exist to remove contaminants. A 5 minute sample workup could increase the signal intensity sufficiently that you are able to obtain neat and meaningfull fragmentation spectra. Good luck!
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Hi,
I am trying to purify a nuclear protein from plant extracts and co-purify any possible interactors using competitive elution as my protein of interest is 3xHA tagged. I confirmed the presence of the protein in the elution fraction by western blot, however when I try to silver stain it, I get nothing. I have tried to modify virtually almost every step in my protocol, yet still nothing on the gel... Any suggestions before I go mad? I need to get it stained before I send it to mass spectroscopy. Thanks!!!
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I think some metalloproteins stain negative with silver stain which may be less visible than a positive stained control
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Hi, 
I have done a hundreds time of denatured Nickel purification. This is my first time encounter this problem. 
I was using 6M GuHCl, 300mM NaCl, 50mM Tris pH 8 to wash the column after loading my lysate. Everything looks fine till the moment I added 250mM of Imidazole in my wash buffer to elute my protein. The Nickel started stripping off the column. I couldn't think of any reason for this. I thought the resin was old so I changed to new resin and same thing happen. There is no metal binding domain in my protein. 
Please help!
Thanks.
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Something must have changed without noticing it. We are dealing with coordination bonds which can only be perturbed by a low pH or the presence of a competing molecule. I would prepare new solutions with new reagents. 
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Hi
I tested the stability of catalase during freeze-drying.
I have found through FT-IR that the amide 2 of catalase has damaged during freeze-drying.
The catalase activity assay result also showed about 50% less than the native catalase.
I want to know the relationship between catalase activity and amide 2.
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I see. Thank you!
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I'm trying to do total protein quantification with Stain-free precast gels and after a lot of reading and many phone calls with Bio-rad, I am confident it's possible to read the gels and membranes with just UV box or an imager with UV capabilities. Has anyone done this before? How have you been able to get good images and measurements? So far, using a G-Box imgaer has given me pretty bad pictures, but I can see the bands so much clearer on just a regular old UV transilluminator box in my lab. Any suggestions for getting better images for normalizing total protein? 
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Hi,
maybe this publication could help:
Btw, you could also "stain" standard SDS-PAGE gels by soaking in TCE after running the gel, but then there is not much of an advantage left compared to other staining methods. The method also works with Trichlor-acetate (only after the run, because TCA can't be added to the gel directly before the run due to its charge).
I guess the imaging system has been optimized in terms of wavelength, exposure time and image processing so if you can afford it, you can save some time. Or spend some time at a regular imager and optimize yourself.
Hope that helps.
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I take the cells trypsinize the cells and wash them with cold PBS two times.
Then i use RIPA lysis buffer according to the pellet and add protease inhibitors 1/20th volume of lysis buffer. Pipet it for mixing and then incubate on ice for 15 mins. then i sonicate it for 3 cycles of 20 seconds each.
Further i centrifuge at 14000g for 30 mins at 4C.
and then take the supernatant and make aliquots.
and i am getting these type of gels.
Thanks 
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Thanks a lot i will follow all this
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Hi,
The spectra of diphenylalanine amide monomer (red) and of dialdehyde crosslinked FF amide dimer (blue) are given below.
What is your interpretation on this spectra?
Which peak(s) belong to amide I, amide II and amide III band?
Which conformational changes occur after cross-linking?
Thank you for your help :)
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Dear Betul:
After studying your spectra - my advise to you: better change the method. In my view FTIR is incapale to solve your problem.
Kind Regards,                              Leonid
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The AcrA protein has purified from the E.Coli using a buffer containing Tris-Hcl, NaCl, PMSF, Triton-X and imidazole
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Thank you, I have not tried earlier. I will consider about that;
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I am trying to design multi-target directed ligands. I have found some similar common ligands for both proteins. When I docked these ligands I have seen the score is almost similar. But these ligands are not that much potent compared to other inhibitors. So I wanted to modify them and achieve some good potency. So can anyone please tell me how to compare the active sites of two different proteins?
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Dear Keith. Thanks for the detailed description. I will see these links and will use it for my studies
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Hi!
I'm performing a DSF assay using a known ligand of the protein I'm working with. I have observed that increasing the concentration of the ligand the melting temperature of the protein tend to slightly decrease, as the fluorescence intensity  emitted by the binding of the SYPRO ORANGE dye to the unfolded protein. Shouldn't I expect the opposite? Could it be possible that the ligand destabilize the protein, instead of stabilizing it? 
Thanks very much for your help
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