Questions related to Protein Biochemistry
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!
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?
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
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.
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.
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?
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.
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?
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?
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,
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,
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.
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.
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?
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?
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.
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!
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!
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?
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!
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).
any technique which can be used to tell the functional and bioactive properties of keratin
any other theoretical reason
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).
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?
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?
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:
.TPP (thiamine pyrophosphate)
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!
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.
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?
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.
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
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 . 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.
 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
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?
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
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.
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.
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 ?
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.
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.
I am looking for methods that can help me in identifying natural small molecules binding with my protein of interest.
Thanks in advance.
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?
Thanks for your valuable suggestions.
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?
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?
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?
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 ?
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.
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.
I have tried to dimerize this protein without success, both proteins are purified
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?
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?
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
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.
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.
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.
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.
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
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?
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?
can acidic pH and guanidinium hydrochloride and high temprature cleavage the protein intermolecular disulfide bond ?
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?
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?
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?
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.
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.
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