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Currently, I am facing a problem with my beta amyloid peptide (1-42) not showing any neurotoxic effect to my SH-SY5Y cells (P24).
Is there anything wrong with the protocol I have followed. Below is the brief protocol I have followed.
I have screened the for neurotoxicity effect at 10, 30, 50, 100 uM beta amyloid concentration.
Day 1:
  • Seed 36 000 cells per well
Day 2:
  • Prepare beta amyloid peptide stock (10mM) fresh where 1 mg is dissolved in 22.2 uL of DMSO (100%)
  • Prepare 100uM beta amyloid peptide: Aliquot 9.6 uL of 10 mM peptide stock into 950.4 uL of complete DMEM (10% FBS+5%penstrep).
  • Serial dilute 100uM peptide solution using 1% DMSO to prepare 50, 30 and 10 uM peptide concentration
  • Aliquot 100uL of each peptide solution (in 1% DMSO) into each treatment well.
Day 3:
  • Aspirate, aliquot 100uL MTT (0.5 mg/mL), wait 4 hours and aliquot 100uL 100% DMSO. Read absorbance at 540 nm with reference wavelength 690 nm.
Do advice me on how i should prepare the beta amyloid peptide to assure neurotoxic effect is seen.
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Thank you for the reply.
Do you have a recommended protocol on aggregating the beta amyloid (1-42) peptide beforehand?
May I know if you have tried out the neurotoxic effect of beta amyloid (1-42) on SH-SY5Y cells that are undifferentiated cells? Mostly i have been seeing articles that are for differentiated SH-SY5Y cells instead.
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Dear scientists,
I got a set of around 4000 protein ids from a proteomic experiment and I would like to globally analyse if the particular groups of proteins in my experiment are significantly more hydrophobic and/or aggregation-prone compared with other groups. I am looking for an R programming library or a web tool that will enable me to obtain some quantitative value for hydrophobicity per protein for my sets. One thing I may do is to just simply calculate the sequence length adjusted number of hydrophobic amino acids C, L, V, I, M, F, W but this seems to be a little naive and I am not sure about the biological relevance of such a simple calculation not taking into account the whole structural aspects of the sequences...I would be glad for ideas on any smarter approaches...please help
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Basically, what you want to do is to determine the amino acid composition for each sequence (e.g. in this R package https://cran.r-project.org/web/packages/protr/vignettes/protr.html using extractAAC()) and multiply the number of times a given amino acid occurs with its hydrophobicity index in your chosen scale (see https://web.expasy.org/protscale/ for different scales) and sum up the values. However, in my experience, hydrophobicity is a poor predictor of the aggregation propensity of folded proteins, as aggregation is frequently linked to imperfect folding rather than to the association of properly folded molecules.
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Can I use protein with glycerol (added to prevent protein aggregation, and maintain stability during -80 storage) in mice (intraperitoneal) to study anti-inflammatory properties of the protein?
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Hi Komal, did you end up using glycerol, what percentage and what was the outcome? Best wishes,
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The protein sample was analyzed after dropping. The buffer had NaCl in it. I just know that something that looks black might be air bubbles. But here these particles are elongated. So I couldn't identify it. I would be grateful if anyone here would suggest what these are.
Thank you.
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Allan Philippe
Thank you for your suggestion. I have attached here two pictures after dropping and then degassing the same sample. The proteins had both fibrillar and amorphous aggregates after degassing. But still doubtful why proteins could look black in the images.
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Hi I am trying to express my protein R in tobacco leaves and do a CoIP, but I can't detect the protein R in CoIP input.
I took the same tissue and added 8M Urea plus LDS protein buffer, protein R shows as a distinct band at the correct size. But when I grind the tissue and put in CoIP lysis buffer, after centrifugation, I put the input in LDS, I couldn't detect my protein, I only see a smear.
Does anyone know what went wrong?
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ok, we´re working with the Plant Protease Inhibitor cocktail from Sigma (P9599).
This works also fine.
Best and good luck furthermore!
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Hi All,
Recently, I purified 8 proteins and stored at -20 with the elution buffer containing 20 mM Tris, 250 mM NaCl and 200 mM Imidazole. I autoclaved the buffer, so I do not know whether the buffer is still fine or not. I did not add triton X and glycerol. My protein concentration was around 1 to5 mg/ml. I stored my proteins at -20 and next day I took them out to thaw. Unfortunately, All my proteins formed aggregate. I could see the white clump. So, I span down and re quantify. protein concentration was reduced by 90%. Could you please share your opinion on how to prevent aggregation from freeze-thaw? If my protein is stable at 4C for 10-15 days, should I still need to store it at -20 or -80C? I am planning to elute my protein in Sodium phosphate buffer with 0.1% Triton-X and 5% glycerol. Should I also use NP40, PEG6000 along with Triton X.
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Hello Abir, there is no general rule, some proteins like salt some other don't, so you have to try both. Make sure also that the isoelectric point of your protein is not close to the pH you use, and if it is, change the pH of your buffer to at least 1 pH unit away from it, and be careful that the pH of a Tris buffer is quite temperature dependent. Alternatively, imidazole can promote aggregation in some instances, so you can try to keep the salt but remove the imidazole (sometimes diluting the sample with elution buffer without imidazole just after it exits the column to something like 50 mM imidazole does the trick). Indeed it is possible that aggregation begins even before freezing and that freezing-thawing just makes it worse. Finally, try to flash-freeze as much as possible (in liquid nitrogen or ethanol+dry ice ), slow freezing can damage proteins.
Olivier
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I am trying to do a kinetics characterisation of AB-40 and AB-42 (they are recombinant peptides produced in the laboratory) but it seems that the samples ar alredy aggregated when I try to do the kinetics mesurement. This peptids, after the purification procedure, are storaged at -20°C and in a solution with a pH of at least 8. I was trying to disaggregate using sonication but it doen't work so well, i was wondering if there is another method I can use for the sample preparation.
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You refer to amyloid beta peptides that are highly hydrophobic peptides. I came across the following paper that might be useful to you:
Broersen, K., Jonckheere, W., Rozenski, J., Vandersteen, A., Pauwels, K., Pastore, A., ... & Schymkowitz, J. (2011). A standardized and biocompatible preparation of aggregate-free amyloid beta peptide for biophysical and biological studies of Alzheimer's disease. Protein Engineering, Design & Selection, 24(9), 743-750.
If you have a powder form of the peptide you could ‘simply’ use DMSO or TFE (after TFA treatment) see for example:
Killian, J. A., Keller, R. C. A., Struyve, M., De Kroon, A. I. P. M., Tommassen, J., & De Kruijff, B. (1990). Tryptophan fluorescence study on the interaction of the signal peptide of the Escherichia coli outer membrane protein PhoE with model membranes. Biochemistry, 29(35), 8131-8137.
Where a highly hydrophobic signal peptide was used.
Hope this helps you a bit further.
Best regards.
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Dear All, I am purifying my protein (theoretical PI = 8.9) in a buffer of pH =7.1. till Ni-elution I kept the DTT concentration (1mM). Usually, after elution people used to keep DTT concentration at 5mM. But in my until Ni-elution the protein look absolutely fine. But the moment I add DTT (5mM) thread like aggregate is formed. when I run those aggregate, it shows up as my protein. What could be the possibilities that DTT is making it precipitated. As I know DTT is used to solubilize protein as reducing disulfide bonds. I will appreciate your suggestions.
Thank you
With kind regards
Prem
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Dear Prem
the effect of dtt and other reducing agent is not ever positive, it depends from the role of The cisteines, if the cisteines are involved in The formation of a structural s-s ( as for example happen in Cu,Zn sod1) the addiction of Dtt may induce protein unfolding and aggregation it precipitation. On the contrary if your protein contain just 1 cystene or the cisteines do not have any structural role but for examples are involved in The binding of a metal (as happen in Sco1) Then the addiction of Dtt can play a positive role.
you can find a more detailed explanation of it on my blog:ProteoCool (https://proteocool.blogspot.com/) in the presentation:
ProteoCool n°16
(3 Common mistakes in buffer preparation for protein ) at page 4
good luck
Manuele
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Am using a 160mM HEPES buffer and diluting serum by 10x and 200x. Found this [ ] works with blood.
pH of buffer is 7.4 and serum was kept at either 4C or RT.
Disperant T was altered from 4C to 37C, allowing for a 2 min equilibration time.
Conducivity lowered with decreasing Temp: 37 (4.74) to 4 (1.94)
Used a voltage of 50V.
4C: -4 AVG ZP
25C: -7.5
37C: -10
For every measure I got "count rate varies- sample not stable clean"
Can anyone point out some literature sources or provide technical support?
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Daniel Villarreal When you say serum, does that imply that you have multiple protein species present? If so, zeta potentials reported by the instrument will be of very little use because it measures an average value (if you have a Zetasize Nano or newer running in monomodal mode) or utter gibberish if running in the mode that gives you a distribution.
Please post an example .dta file so I can look at the various diagnostic information.
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I've been testing my A-beta 25-35 samples with ThT assay on a black 96-well plate to monitor its aggregation state, the fluorescence in 3 replicate wells differ dramatically unless I pipette the wells right before assay. 
I've read about shaking the plate before each ThT assay, but I'm not sure in what ways this method helps, and how can I shake a 96-well plate sufficiently without spilling the contents?
Thanks!
Lin
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Hello!
Shaking delivers extra energy to your system - it can shorten the lag time
Best
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I am using a bacterial protein and its get aggregated. I get a clear band of appropriate size in SDS PAGE but unfortunately it forms smear pattern in NATIVE PAGE. Also i try to check in TEM and found several aggregates.
Proteins are purified using Flag Affinity Purification, followed by gel purification.
pI of my protein is 7.28 and buffer I use is TBS of pH 7.4 (at 4C)
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The chromatogram from your gel filtration step is a source of information regarding protein stability and a well folded protein, many scientists focus too much of SDS-PAGE gel, in fact often just using SDS-PAGE for selecting fractions instead of using both.
GF buffer selection is a part of good gel filtration practise as the protein will be in that buffer for a considerable amount of time in this environment.
I don't like Tris as it is not really a good buffer. For GF I generally use 10-25mM HEPES 7.5 (in your case I would use pH 8.0), with 125-500mM NaCl, 5% glycerol and 0.5mM TCEP (non-thiol reducing agent with v. good stability).
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I have expressed my 70 kD protein (pI 6.7) in BL21 E.coli cells and purified it using TALON metal affinity resin (cobalt-based resin). My elution buffer is 20 mM Hepes Ph 7.6, 500 mM NaCl, 10 % glycerol, 1mM TCEP, and 150 mM imidazole. My previous experience with this protein had shown that it tends to form some sort of dimers/aggregates thus I added 1 % octyl-glucoside (OG) and 5 mM EDTA to the eluted protein that were in a buffer mentioned above. Although addition of 1% OG was borne fine, upon addition of 5 mM EDTA the solution went cloudy and after 48 hour incubation at 4 degrees there were lots of aggregates in the solution. I spun down the aggregates and tried to resuspend in 8M urea but it wasn’t successful attempt.
I thought addition of EDTA might be beneficial through chelation of any leached cobalt ions. Is it a possibility that chelating leached cobalt ions also precipitated my protein?
Or chelation of useful divalent ions knocked my proteins out and precipitated them?
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I know this question is quite old already, but since I found It as the top results on my google search I thought I might help any future visitors.
It is true that EDTA can cause problems for his tagged proteins, the stability of a quality Ni-NTA agarose resin is about 1 mM here. Every concentration above that will have severse consequences. When working with NTA or IDA based resins at least.
The INDIGO ligand has an EDTA stability of 20mM, so 20 times the concentration of quality Ni-NTA. That should be used here.
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We are working on the fibrillation of lysozyme. Given the acidic condition (pH 2) of the protein solution (glycine buffer) for fibrillation, what should be buffer and its pH for ThT stock solution?
Should the buffers of protein and ThT solutions be identical?
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I have never worked at such low pH with ThT. The best way would be to put ThT in water or in the same pH of your buffer but the efficiency of the absorption and emission will change with respect to physiological pH. I would suggest you to give a look at this paper (DOI 10.1007/s00249-015-1019-8 ) in which ThT was tested in a wide pH range.
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REFERENCE 1: Sanjay Kalra, Jubbin Jagan Jacob, and Yashdeep Gupta. Newer antidiabetic drugs and calorie restriction mimicry. Indian J Endocrinol Metab. 2016 Jan-Feb; 20(1): 142–146.
De-acceleration of aging and delayed development of age-related morbidity accompanies the restriction of calories (without malnutrition) in laboratory mice, nematodes, yeast, fish, and dogs. Recent results from long-term longitudinal studies conducted on primates have suggested longevity benefits of a 30% restriction of calories in rhesus monkeys as well. Among calorie restricted rhesus monkeys one of the mechanisms for the improvement in lifespan was the reduction in the development of glucose intolerance and cardiovascular disease. Although there are no comparable human studies, it is likely that metabolic and longevity benefits will accompany a reduction in calories in humans as well. However, considering the difficulties in getting healthy adults to limit food intake science has focused on understanding the biochemical processes that accompany calorie restriction (CR) to formulate drugs that would mimic the effects of CR without the need to actually restrict calories. Drugs in this emerging therapeutic field are called CR mimetics. Some of the currently used anti-diabetic agents may have some CR mimetic like effects. This review focuses on the CR mimetic properties of the currently available anti-diabetic agents.
REFERENCE 2: Appetite Regulation and the Peripheral Sink Amyloid beta Clearance Pathway in Diabetes and Alzheimer’s Disease. Top 10 Commentaries in Alzheimer’s Disease (e-book). 2019;2:1-11. www.avidscience.com.
REFERENCE 3: Diet, Drug and Inhibitor Therapy Prevent Toxic Protein Aggregation in Various Species. Acta Scientific Nutritional Health. 2(8);2018:01-03.
REFERENCE 4: Calorie Sensitive Anti-Aging Gene Regulates Hepatic Amyloid Beta Clearance in Diabetes and Neurodegenerative Diseases”. EC Nutrition ECO.01 (2017): 30-32.
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Please check
1. Alessandra Stacchiotti, Gaia Favero and Rita Rezzani .Resveratrol and SIRT1 Activators for the Treatment of Aging and Age-Related Diseases.2018. DOI: 10.5772/intechopen.78977
2. Alberto Zullo. Emanuela Simone , Maddalena Grimaldi, Vincenzina Musto and Francesco Paolo Mancini. Sirtuins as Mediator of the Anti-Ageing Effects of Calorie Restriction in Skeletal and Cardiac Muscle . Int. J. Mol. Sci. 2018, 19, 928.
3. Niria Treviño-Saldaña and Gerardo García-Rivas, “Regulation of Sirtuin-Mediated Protein Deacetylation by Cardioprotective Phytochemicals,” Oxidative Medicine and Cellular Longevity, vol. 2017, Article ID 1750306, 16 pages, 2017. https://doi.org/10.1155/2017/1750306.
4. David G. Le Couteur Andrew J. McLachlan Ronald J. Quinn Stephen J. SimpsonRafael de Cabo.Aging Biology and Novel Targets for Drug Discovery.The Journals of Gerontology: Series A, Volume 67A, Issue 2, February 2012, Pages 168–174,https://doi.org/10.1093/gerona/glr095
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I'm looking at a protein of approximately 140-150 kDa which is larger than we have ever looked at in our lab so I needed to adjust some steps in our Western blot protocol. 
From suggestions I found online I used 6% gels and started transferring overnight (16h) at 20V instead of 1h @ 300mA like we do for other smaller proteins.  The transfer seems successful and I'm getting bands but I'm getting these huge bands all the way on the top of my running gel at about 260kDa.  I see them even after staining the membrane with fast green (which we do as a loading control for total protein) so its not an antibody issue.  I have never had this before with any of my other proteins, therefor I think it must be caused by the different methods used.
Does anyone know if under these new conditions (6% gels + transfer overnight) it might be more likely to get protein aggregation in the top of the gel for some reason?
Thanks
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Try going for higher percentage of beta-mercaptoethanol in loading dye if you think protein aggregation could be an issue.
Best of luck
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I am planing for cryo-em of a AAA+ ATPase (natively hexamer of 360 Kda)
properties> forms multimer of hexamer at concentration above 2mg/ml and below Nacl 500mM.
Also show denaturation on freezing(required to freez for transporting to cryo-EM facility)
Earlier I did SAXS of this protein and got Aggregated profile from low to high concentration. buffering condition was 150Mm Nacl, 5% Glycerol ,10mM Arginine, 4mM bME, 25 mM HEPES 7.5pH.
i want to know Whether this buffering condition suits cryo-EM requirement?
If SAXS is showing Aggregated profile, is it possible get single hexamer images from cryo-EM in same buffering condition(given above) ?
what are the precaution required to take care during sample prepration?
this will be my first time on cryo-EM.
All the assistance and suggestion are welcome
Regards
Ketul Saharan.
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Glycerol can cause problems for electron microscopy, but 5% may still be ok. If you see that your sample starts to look strange with bubles this is likely due to the glycerol, which is sensitive to radiation. An idea could be to prepare at least some sample that does not contain it or to be prepared to dialyse some sample before freezing.
Spinning down the sample may also remove some of the aggregates. A good way to optimize your sample would be to start with negative stain. This is much faster so you can try a few things before doing cryo. It will already show if the aggregates are a problem, or whether you have still enough single protein to proceed. It will also alow you to optimize the concentration of protein.
Sample preparation for cryo-EM is mostly trial and error, so the best advice is basically just to have a look and then go from there.
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As a chemist, I am working on hyaluronic acid (HA) hydrogels, which are DNA-modified. DNA is attached to HA via Thiol-Michael. However, due to the polyanionic character of HA, DNA forms aggregate during hydrogelation. I used 15 mM Phosphate Buffer and also tried 15 mM Phosphate Buffer 300 mM NaCl, but both didn't work.
Gernerally, what could I do to rule out this aggregation issues? Are there any additives that could be added?
Thanks for your help!
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Of course, happy to help!
1. I think that freeze/thaw is fine, wouldn't hurt to add a little DTT to the storage solution though. Thiols aren't so reactive that a bit of casual exposure to the air will ruin them, it's just in the PCR reaction where it's being heated to 95 repeatedly that some irreversible oxidation to the sulfone/sulfoxide/sulfate could be an issue.
2. I don't think you need a protected primer, I have used 5'-thiol-C6/maleimide chemistry a few times without difficulty. I don't think it's worth the extra hassle of deprotecting your finished PCR product. Biggest thing is just to remember that even though this is a biochemical reaction we should always take the same precautions we would when doing organic chemistry in terms of using inert atmosphere where appropriate etc.
3. You only need about 5 - 10 mM of a cation to keep the DNA in the double stranded native state, 50 mM buffer is a bit extra but will make sure the pH is steady. 250 mM salt (buffer + NaCl) is quite high. Theoretically it shouldn't interfere but it's always best to keep things as simple as possible. Higher salt neutralizes the negative charge of the DNA and can cause it to form aggregates in solution and do other weird things. Again, shouldn't interfere but why take the risk?
4. Cool method for determining success! It would be difficult to see even a 40k addition by agarose gel electrophoresis, since that would correspond to only roughly a 60bp difference, but if you run a 5% acrylamide/TBE gel you should be able to see the modification, perhaps even your 10k PEG moiety. However, these gels take a bit of practice to get them running perfectly.
Best of luck!
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Hi all,
We have HiLoad Superdex 200 16/60 Gel Filtration column in our lab, which have some 'Sticky Proteins' and protein aggregates bound to the column. As such, I wish to perform some rigorous cleaning of the column to get rid get rid of these unwanted debris on the column?   
I have done some reading from the column manufacturer manual (GE) and they have recommended 30% isopropanol and 70% of ethanol together with high denaturing agents (Ie. 6M  Guanidine Hydrochloride/8m Urea).
Based on your  experience and opinion, what would be the best approach/cleaning solutions.
Thanks.
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Hi,
first with 30% isopropanol 1 CV, wash with 1 CV H2O
- Wash with 0.5 M NaOH 1CV, wash with 2 CV H2O
- additional cleaning using 500 mM acetic acid can be useful. Wash with 2 CV H2O
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I am going to overexpress the alpha-synuclein in the SH-SY5Y cells in order to investigate some drugs on the inhibition of aggregation and the mechanisms that inspire such cells go under apoptosis. The question is how I could detect the protein aggregation in the overexpressed cells. Is it possible to detect Lewy body-like inclusions in the cells? In fact I am going to distinguish the differences of the alpha-synuclein aggregation between the overexpressed cells and such cells while treating by some small molecules.
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Hi Farhang,
I'm not sure if you're still looking into this but here are protocols we use for thioflavin T and cell culture assays to detect alpha synuclein aggregation:
Good luck!
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ABSTRACT
Genomic medicine treatment of diabetics is critical in the current global diabetes epidemic to prevent the expected diabetes pandemic predicted to occur by the year 2035. The major concern with Type 3 diabetes in the global population is related to a defective SCN with relevance to uncontrolled peripheral glucose levels and endocrine autoimmune disease. Appetite regulation and genomic medicine are critical to Sirt 1’s regulation of the MHC genes with relevance to maintenance of immune recognition and endocrine hormone treatment of mitophagy. In the developing world the major concern for a diabetes pandemic is mitophagy and will require diets with Sirt 1 activators to prevent Type 3 diabetes, endocrine autoimmunity and mitochondrial disease.
RELEVANT REFERENCES:
1. Martins IJ. Genomic Medicine and Endocrine Autoimmunity as Key to Mitochondrial Disease. Glob J Endocrinol Metab .2(2). GJEM.000534.2018
2. Martins IJ. Biotherapy and the Immune System in Ageing Science. Acta Scientific Nutritional Health 2.4 (2018): 29-31.
3. Martins IJ. Appetite Control and Biotherapy in the Management of Autoimmune Induced Global Chronic Diseases. J Clin Immunol Res. 2018; 2(1): 1-4.
4. Martins IJ. Heat Shock Gene Inactivation and Protein Aggregation with Links to Chronic Diseases. Diseases. 2018, 6;39:1-5.
5. Martins IJ. Autoimmune disease and mitochondrial dysfunction in chronic diseases. Res Chron Dis (2017) 1(1).
6. Martins IJ. Regulation of Core Body Temperature and the Immune System Determines Species Longevity. Curr Updates Gerontol. (2017) 1: 6.1
7. Martins IJ. Anti-Aging Gene linked to Appetite Regulation Determines Longevity in Humans and Animals. International Journal of Aging Research. 2018,1(6): 1-4.
8. Martins IJ. Genomic medicine and acute cardiovascular disease progression in diabetes. Res Chron Dis (2018) 2(1), 001–003.
9. Martins IJ. Genomic Medicine and Acute Cardiovascular Disease Progression in Diabetes. International Journal of Medical Studies. 2018;3(1): 124-130.
10. Martins IJ. Electroconvulsive Therapy and Heat Shock Gene Inactivation in Neurodegenerative Diseases. Ann Neurodegener Dis. 2018, 3(1): 1028.
11. Martins, I.J. (2018) Indian Spices and Biotherapeutics in Health
and Chronic Disease. Health, 10, 374-380. WITH AND WITHOUT INDIAN SPICES
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IT is a beautiful question. I will myself be interested in the answer.
As far as I think, even if some therapy is curative by how can you regain the lost beta cell mass?
A question worth pondering upon!
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For my project I need to have purified HEWL fibrils without oligomers and monomers and vice versa. I couldn't find any useful protocol for this separation, I also wanna know if there is any protocol to force all HEWL monomers get into fibril form without any residual monomer and oligomer.
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What should we do, if we don't get any pellet formed after centrifuging?
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I am working at industrial scale enzyme production. Where I found precipitation after filtration and concentration. Sometimes during concentration itself material becomes turbid. Is there solution to get rid of this?
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Thanks Mr.Omar for your prompt support.
May I know the processes which you are suggesting to increase solubility of protein?
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I have to ship a batch of purified proteins at ambient temperature to a collaborator. The functional state, activity, and folding of the proteins after shipping are not important as they will be used for proteomics analysis. The proteins have been His-tag purified and precipitated with the methanol-chloroform-water method. I am wondering whether it would be better to ship the proteins as precipitates or freeze-dry the precipitate before shipment. Maybe the protein will be less stable as a precipitate compared to as freeze-dried during shipment, thus causing degradation and loss of material. On the other hand, re-solubilisation of freeze-dried proteins could be challenging, which might also lead to loss of material from incomplete re-solubilisation.
I know that each protein could behave differently but I would very much appreciate any comments on whether in general shipment as precipitate or as freeze-dried at ambient temperature could be better, i.e. lower degradation and loss of material?
Thanks a lot in advance!
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I have had mixed results reconstituting freeze dried proteins from vendors, and consider it something of an art. Protein left in an "elegant cake" has a lot of surface area and takes to rehydration well, while protein that is packed into a dense film doesn't solubilize well. The company (Hypermol) that provides us with readily soluble protein mentioned to me using unnamed (read: proprietary formulation) sugar in their formulation, and I have worked with protein formulated with trehalose (and surfactants and whatnot) in the past that has performed well under lyophilization/reconstitution.
I know that doesn't take you all the way to a solution to your problem, but hopefully it gets you a step in the right direction. Good luck!
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Hi, everyone,
I am interested in the M13 bacteriophage, but similar species might be relevant.
Is is possible that, an over-supply of one phage protein (more copies than overall phage replication/exit can keep up with) could result in that phage protein clumping together into aggregates?
My idea is that, a mutant phage isn't fully assembling, and then. the non-assembled proteins might just gather together, due to being very hydrophobic(?)
From some recent reading, it seems that, certain mutant phages with altered protein expression will kill the host (E. coli).
I am wondering if this is caused by surplus phage proteins, sort of clogging up the host(?)
Thanks for any ideas...
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The assembly capsid proteins shall anchor on the ecoli surface. Simply added ectopic proteins should not change the assembl. Like Anne said, the extra coat protein will be stay within ecoli, or in the pe.
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For amyloid-beta 25-35, many papers report using distilled deionized water to dissolve, then incubation at 37 degree for 3 days to make it aggregation. I already tried, the prestoblue results and hochest stain showed there was no cell viability difference between control and experimental group after incubation with different concentration peptide(5uM-50uM) for 24 or 48 hours.
For amyloid-beta 1-42, I saw one paper (Feng and Zhang, 2004,J.Pineal Res.) used sterile water to make it reconstituted (400uM) at 37 degree for 72h. I tried, but the amyloid-beta 1-42 did not dissolve in water even at 200uM concentration. And also have no toxic to neuroblastoma cells. Does anyone have experience on preparing these two kinds of peptide? Thanks so much!
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Dear Thomas
How long can you use the stock A25-35 after your incubation?
Do you always need to use freshly?
Thank you in advance!
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Hello,
I have been recently putting a lot of effort in finding any information about amyloid fibril formation in mabs (or mab fragments, or other protein therapeutics) in current scientific articles. Unfortunatelly, with no success. On the other hand, I have encountered countless number of articles describing fibril formation in peptide therapeutics (e.g. insulin, glucagon or calcitionin).
1. Could large protein therapeutics form amyloid fibrils?
2. Is there any clear line above what size (or amino-acid content) proteins do not form amyloid fibrils?
3. Have you encountered any articles concerning amyloid fibril formation in protein therapeutics?
Best regards,
Dorian
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Hi Dorian, proteins forming amyloid fibers are reported quite frequently. These are usually small proteins or peptides that can assume various conformations including fibril structures.
Antibodies have a very constrained conformation so it is unlikely that they will form fibril structures.
But to be considered as classical "amyloids", protein aggregates have to show binding of thioflavin T with accompanying increase in fluorescence.
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I am trying to measure the Zeta-potential and protein mobility of LDH A (332aa, 33kDa). Since we have a very limited amount of the protein, we decided to use the diffusion barrier technique with 0.5mg/ml LDH A suspension. We encounter a big problem with protein aggregation, thus getting unreliable results. I would like to measure at pH 4-10, but even test measurements at pH 7 (pI for LDH A is 8.1) would fail.
The software would give values for zeta-potential and mobility, but except for the size distribution, no graphs would be generated and l would constantly get the message that data quality is poor due to aggregation happening not only before the measurement, but during the measurement as well (shown by a steadily increasing PDI). I would always use the Auto mode and Monomodal settings, but the Zetasizer would occasionally even abort the measurement.
I tried the same sample preparation and measuring procedure with BSA, but the aggregation problem persists. Even after testing in different buffers (Tris, HEPES, PBS) and different agents to prevent aggregation (Tween20, TritonX100, sucrose, glycerol, L-arginine, guanidine hydrochloride protocol), the majority of the sample would still stay aggregated.
The question is, is there a more reliable way to prevent protein aggregation before and during measurement? I don’t expect a perfectly monodisperse sample but something that would get me closer to a more acceptable PDI.
Or, if possible, is there a way to bypass the results of the aggregated protein (even though it is the majority of it) and only use the values for single protein peaks? What I mean by that is that is, out of two peaks that appear in the intensity graph (one for single protein, one for aggregated protein), can the second peak be deleted out of the equation and only to one remain for calculations, no matter the intensity?
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Hi Anela,
Starting with the most alarming points you raise, what error message did you get when the Zetasizer aborted the measurement? The instrument will do some optimisation checks at the start of a measurement and there are certain conditions it checks for before starting the measurement, so more information on the error would be helpful to enable a solution to be suggested.
What version of the Zetasizer are you using? The Zetasizer Nano ZSP has a higher power laser than the other variants meaning that is has better capability to measure proteins and other low scattering samples.
Also, your concentration may be a little low. The specification point for zeta potential measurements based on lysozyme (14kDa) is 1mg/ml. You should expect to be able to measure lower than that for your sample given that it is larger, but when using the diffusion barrier method there may be some degree of dilution as the sample diffuses over time, making measurements more challenging with a low volume of low concentration sample.
Are you running zeta potential measurements or protein mobility?
Both zeta potential and mobility are reported for both measurement types but the processing is different between them.
In a protein mobility measurement, the data is analysed as the measurement progresses and data that is expected to be attributed to the detection of stray aggregates is omitted from the final result, so even if some aggregation does occur, the results can still be reliable.
Additionally, the protein mobility measurement performs a size measurement before and after the mobility measurements to check on the state of the sample. If this shows aggregation, it does not indicate that the measured mobility is invalid (as per the processing discussed above), but that the sample is not suitable for repeat measurement.
Mono-modal measurements will not produce a zeta potential distribution as this data is calculated from the Slow Field Reversal part of an applied waveform in a General purpose measurement. This is omitted from a mono-modal measurement for samples that are either fragile or suspended in a high concentration solution in order to reduce the possible effects of Joule heating.
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ANS dye which is popular to study exposed hydrophobic patches in a protein structure, i have added it with hen egg white lysozyme but after few hours it gets settled down, what could be the reason as i suspect its not able to interact with protein.
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What do you mean "it settled down?" If you see black particles i guess you are using actual ANS and not its salt. Try using ANS amonium salt and disolve it in PBS buffer (ph 7.4) it should work.
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I am working on an integral membrane protein. I purified this by NI-NTA resin successfully. But, when I concentrated this protein by using centrifuge at 3000rpm, 10min and 3min, this protein has been aggregated.
I used an 100K cut-off membrane, and solubilized this protein by using the buffer containing 1% DDM, 0.2% CHS, 20mM HEPES pH 7.5, 200mM NaCl.
I don't know what happened.
I thought it is because of eluted buffer containing imidazole, but even though I removed the imidazole by dialysis, It has been aggregated and precipitated.
Is this because of behavior itself? or do I need the detergent screening during concentration?
In detergent screening in just fSEC, I got the best peak shape and height in detergent and Fos cholin.
Can anyone have suggestions?
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Thank you, Omar Gonzalez-Ortega !
I will try the more test like your suggestion.
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I purified my protein from inclusion body by solubilizing it in 0.1%  N-lauroylsarcosine -PBS. I wanted to reduce the amount of N-lauroylsarcosine, I conducted dialysis in PBS overnight. Next day I found that the protein was aggregated. How can I prevent protein aggregation?
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I would like thank all of you for the valuable suggestions.
I try to change salt concentration ranging from 300 to 500nM as suggested by Ali and under several PH as suggested by Robin. Also I tried PEG 400, 6000 and 8000 with glycin or glycerol.
Finally< Conducted a cytotoxicity test of N-lauroylsarcosine on several cell lines (adherent cells) in order to know the minimal concentration of N-Lauroylsarcosine that can exhibit a toxicity. Then my protein is now soluble in that minimal concentration (between 0.01-0.02%).
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I have tried to generate cellular systems showing Tau protein aggregation induced cytotoxicity in various mammalian cell systems including HEK, CHO-K1, and neuroblastoma cell lines. However, most of my efforts have failed because mammalian cells overexpressing tau proteins showed quite normal proliferations comparing with control cells. Does anybody know how to induce cell death by tau aggregation induction?
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As many suggested above, mere overexpression of Tau in cells will not cause apoptosis. But it could cause some other toxic effects which one need to focus on.  If you would like to study only the apoptotic pathway induced by Tau aggregation in cells, one should use mutant Tau, such as
If you would like to study only the apoptotic pathway induced by Tau aggregation in cells, the following criteria must be met to my knowledge.
1.one should use mutant Tau, such as delka K280, P301L etc. 
2. The fulllength Tau even with the mutation does not form ThS positive aggregates (which are considered as pathological aggregates - presume to contain filaments). Therefore one should use the repeat domain of Tau alone to induce the aggregation inside the cells. In that
In that case, one could definitely observe ThS positive aggregates, cell death and other toxic effects.
3. please find the below the link for the article where you might get the information you want.
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I'm working on a protein with 13 cysteines among 500 amino acids. It aggregated easily, and I want to mutant cysteine(s) to alleviate the situation. How to choose the cysteine that need to be mutant and how to evaluate the result?thank you!
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What information do you have on your protein - do you have a structure, a homology model? The first thing you want to investigate is whether any of the cysteines form disulfide bonds in the native protein - you do not want to mutate these, as they may contribute to the stability of the folded protein. (Check annotations in uniprot http://www.expasy.org for your protein and murine and human homologs and do a blast search of the PDB http://www.rcsb.org to look for structural information). For the unpaired cysteins, you would like to make an educated guess whether they are buried in a hydrophobic environment - in this case, a mutation to Ala (slightly too small) or Val (slightly too large) are the best replacement. For solvent exposed cysteines, Ser would be the isosteric replacement, Ala and Thr might be viable alternatives. 
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I am measuring p53 aggregation using protocols found in journals and have never done the experiment before not has anyone here.
Papers that I am copying used 1-10uM p53 core domain with 5-120uM ThT. I am aggregating by incubation at 37 C for 0 - 60 minutes while measuring fluorescence every 5 minutes. I am exciting at 440nm and measuring 480nm.
Like protocols that I read online, I mix 1mg/ml ThT covered for 1 hour, then adding to my protein. I am also measuring samples without protein for reference.
My readings are very high (over 100,000 units) and decrease over time, which is the opposite of what I am expecting.
What kinds of things could I be doing wrong?
Thanks,
Karen
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Hi Karen,
It might take a while to establish suitable parameters for aggregation assays. Please consider the following things might be relevant
  • aggregation time (6-72 hours for a lot of proteins)
  • protein concentration (1-100 µM) compare to signal/ concentration relation in Fig 6B of linked article)
  • use not more than equimolar ThT
  • optimized excitation (420-440nm) and emission (480-520nm) Some proteins prefer different parameters)
  • agitation like shaking every 15 min before aggregation
  • pH (could be between pH 2-10)
  • salt concentrations NaCl, CaCl2, KCl
  • mild denaturation with urea
  • reducing environment (e.g.DTT)
  • if you have access to a plate reader try several buffer conditions at once, for my first assay I just grabbed several buffers we had available in the lab
  • check the gain settings in the fluorescence reader
FYI: You can mix ThT solution in advanced, filter it through a 0.2 µm syringe filter the solution a 500µM solution can be stored at 4°C in the fridge for several days (please check also your ThT dilution was correct.)
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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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I would like to get BSA proteins aggregates by either centrifuging or heating to size 300 nm or more.
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I would like to get denatured aggregates of BSA , How to do it thermally?
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Hi All, 
I am working with a bifunctional protein from an unculturable bacteria. Currently the protein is His-Tagged with the Trigger Factor Tag. I cloned the gene into a suite of vectors which included a GFP tag, MBP tag and SUMO tag among others but the protein was only soluble with the TF tag. 
Upon biophysical characterization of the protein by analytical ultracentrifugation and differential light scattering,it appears the protein exists as a solube aggregate and elutes off of a S200 10/300 column before the void volume and does not concentrate when using a Vivaspin filter. The protein does not precipitate out of solution.
All buffers are prepared fresh and include 1mM TCEP. 
Expression conditions such as IPTG concentration, additives such as sorbitol and arginine have been added to the growth media as well as buffers to help prevent aggregation. 
I am currently trialing variations in pH, then salt concentration and then the addition of Urea along with various detergents. 
Does anyone have any experience with this? Your help/suggestions would be much appreciated. 
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From your description, it seems that the protein aggregation you observe is happening during or after purification. In this case you can change a few things in your lysis, purification and storage buffers:
1. If you know the pI of the protein, use a pH that is ~2 points above or below the pI
2. As mentioned by others, NaCl at 0.3-0.5M sometimes helps prevent aggregation
3. Add stabilising osmolytes such as sucrose, trehalose, glycerol, TMAO etc at high concentration 1-2M.
4. Add stabilising amino acids. Arginine and glutamate, either alone or in equimolar mixtures at 50-500 mM
5. If your protein has co-factors such as metal ions then add these in trace amounts to your buffer. 
6. Change buffer. Sometimes proteins are more stable in for example, phosphate buffer rather than Tris buffer.
7. Change your constructs e.g. move tags from N to C terminus or vice versa. Cleave off tags. Add small spacers eg GGGS between tag and desired protein sequence.
8. Detergents - use non-ionic or zwitterionic detergents at >1.5x CMC. Non-detergent sulphobetanes can also prevent hydrophobic aggregation.
9. Many proteins are more stable and less aggregation prone in a ligand-bound state. If your protein has a ligand, include it in your buffer.
10. Reducing agents such as DTT and TCEP can help prevent aggregation if there are free cysteines around your protein.
As you can see, there's a lot you can do to optimise non-aggregated protein yield and you have to work systematically through the list. A simple assay for determination of aggregation will help greatly. You can reduce the amount of work if you know the structure of your protein and whether there are exposed hydrophobic or charged regions. Basic information such as pI and hydrophobicity index can help to decide what to prioritise. 
Good luck!
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Hello! I'm desperately trying to find clues to the molecular mechanism of prion replication: how do misfolded (scrapie form) PrPs actually catalyze misfolding of the normal cellular PrPs on molecular/biochemical level? I understand that it is not completely cleared out, but I couldn't find any molecular/biochemical hypotheses. Everyone seems to just skip this issue in any paper I could get my hands in the past few days. Also, what is the biochemical reasons for which such saturation with beta-sheets renders PrPsc protease-resistant? Isn't there really any proteases specific enough to degrade high beta-sheet-content proteins? Or does it have to do with relatively rapid formation of amyloid fibers? Or does it have to do with chaperon malfunction (thus potentially mutation in chaperon genes as well)?
Thanks!
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Hi Artur,
You can read some of the recent reviews by the pioneers of this field such as:
1. Stanley Prusiner
2. Adriano Aguzzi
3. John Collinge
4. Jean Manson
etc
you can find a lot of information in their reviews. If you have any issues in getting these reviews, let me know. I can send you.
All the best
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I know that the intenisty of ThT can vary from assay to assay when monitoring amyloid fibrills, but my intensities are well below what I feel they should be. The intensities range from 0-200, instead of closer to 1000. I'm following a ThT protocol from Anaspecs Sensolyte for ABeta I'm using 200mM ThT concentrations to monitor 20-50uM abeta. I'm using 50mM Tris with 150mM NaCl at a pH of 7.2. I'm agiatating for 15s between reads and monitoring for 72hrs. Prior to this, I'm reconsutituing AB with 5mM NaOH add to HFIP treated AB film, no vertexing, incubating at RT for 5 mins followed by sonicating bath for 5 mins then introducing to plate. I am getting the typical sigmodal shape for controls and the other, but the intenisty just isn't there along with what appears to be a noisy signal in comparison to results seen in literature. Any help would be greatly appreciated. 
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4uM is a low concentration of beta amyloid for ThT assays, so I'm not surprised you don't see anything for that long of a time period. That is not to say that 4uM beta amyloid assays are not doable. I would suggest looking at this paper in the link below because even at 20uM without shaking my ThT assays take 72hrs, and with shaking they were previously not reproducible among triplets. I started using this technique of adding borosilicate glass beads to the plate before this paper came out and was able to get very very reproducible results even across multiple batches, and was still able to observe the effects of my variables. So the paper is applicable to beta amyloid.
Prep is also very very important in beta amyloid experiments. I suggest an alkaline prep over HFIP/DMSO (Stein prep), there are tons of papers using either so that is up to your experimental setup and limitations.
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I am about to start working with amyloid beta peptides (1-42 and 1-40). I have purchased lyophilized peptide from sigma. I would like to know if I can store the peptides as solution and which reagent is suitable for dissolving it? Sigma suggests to initially dissolve Abeta 1-42 in NH4OH and Abeta 1-40 in HPLC water and then to dilute in phosphate buffer and PBS respectively. After dilution upto about 1mg/ml concentration, if I store the peptides at -80 degree will they be as monomers for some time? Or else what should ideally be done? Thanks in advance.
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Thanks for your suggestion. I will do that only.
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I'm facing with a problem in protein aggregation analysis. There are different algorithms and bioinformatic tools for predicting protein aggregation. I'm using the command "hydrophobic patches" on Pdb viewer and Aggrescan 3d to try to analyze my protein. As I can see they don't agree each other about the possible Amino acids involve in aggregation. 
Can anyone recommend a reliable tool? Additionally, I'd like to know on which algorithms the detection of hydrophobic patches is based.  
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HI Pietro,
Depends on what you want to do: do you want to know the aggregation potential of your protein in the unfolded state (then use short stretch predictors such as TANGO) or do you want to predict the aggregation in native state (Aggrescan3D is definitely good, but you could also look at our Solubis)? Are you looking for ways to reduce the aggregation of your protein? 
There are indeed quite some methods and in many cases they agree, but not always. Most have been benchmarked against experimental data and certainly cover some aspects.
There are good papers describing each algorithm, so best to start there.
Cheers
Joost
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Recently I've been purifying a 85kD protein from Rosetta. The yield is pretty good and the 260/280 is around 0.7-0.8. However, when I run this sample on FPLC superose 6, it showed very huge aggregation peak which is not consistent with a similar protein purified from insect cells(usually monomer). I used the same purification procedures. Does anyone have similar experiences? I used pET15b vector and used 1mM IPTG to induce at 17C overnight. Should I try to decrease IPTG concentration and decrease induction time? I've attached the SDS-PAGE and FPLC profile with this.
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Hi,
I would expect the host cell protein to be very different, Rosetta vs insect cells (e.coli vs euk.
I believe that you need to look into the purification of your target protein and change
to be effective to remove the host cell proteins from Rosetta. How do you know that it is aggregates that you see in SEC, could it just be host cell proteins that you do not get rid of with your method "optimized" for purification of your target protein from insect cells?
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Can anyone confirm my suspicions that it's NuPAGE MOPS buffer without the SDS? Thanks.
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The running buffer (without SDS) will depend on what type of gel you are using. I don't think the NuPAGE gels are available without SDS. You can get Novex NativePAGE Bis-Tris gels, for which the running buffer and other reagents can be found here:
I couldn't find out what is in the running buffer.
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When I concentrate protein using filter centrifuge tube, there is always protein precipitate out. How can I manage this situation. Is there anything I can do to prevent this process?
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In addition to what was proposed before, you can try to concentrate it over an affinity or an ion exchange column. the main limitation with these two ways will be your matrix binding capacity. Something else that you can try is also to dialyse against  high peg polymer ( like peg 20000)
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We are doing crystalization for a protein, which could exist as rod-like oligomer in cells. We expressed the protein in bacterial and in insect Sf9 cells. After purification procedures, protein was examined in size-exclusion chromatography (SEC) and was found that most(>50%) of protein is polymer/aggregate(?). This protein is easy to form polymer/aggregate(?) during purification manipulation. We don't know if the polymer in SEC is natural oligomer or misfolded aggregate.  
My questions:
1. Is protein monomer in buffer solution required for crystal formation, even for a naturally potential oligomerized protein? 
2. Can protein form oligomer complex normally and correctly in buffer solution?
3. Are there any samples that naturally protein oligomer (?) in buffer solution formed  successfully a crystal oligomer structure?
Thank you for your time.
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In some cases proteins that naturally form fibrils and are very difficult to crystallise can be crystallised as defined monomers by either masking the oligomerization interface with crystallisation chaperones such as antibody fragments, DARPins or other binding partners (e.g.. Tubulin complexes crystallised in the presence of DARPins that prevent microtubule formation), or by introducing mutations in the oligomerization interface.
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I am using 40uM of the tht for the detection of amyloid. How to know this is the optimum concentration of tht for my assay.   
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Try to perform a titration assay by varying the concentration from as low as 10uM to as high as 1 mM and record the ThT fluoroscence in each case by using any of the spectrofluorimeter (LS55 lying in the lab). Identify a particular concentration beyond which the signal does not increase any further. That should be your optimum ThT concentration for your assay!!! Gud Luck!!!
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i am working on lysozyme protein aggregation study (effects of several chemicals on its structure), i need suggestions for the buffer medium i should consider and why?
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PBS may be used for studying lysozyme aggregation. Please go through the article "Fibrillation of hen egg white lysozyme triggers reduction of copper(II)". There are other articles also by the group on the same topic.
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Protein aggregation underlies several neurodegenerative diseases. The recent evidence suggests that in fact aggregated protein in the living cell can lead to cell death. I would like to know your ideas about the mechanism of this toxicity.
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Dear Laleh,
As a Bachelor student I have worked on poly-Q protein aggregation that is known in Huntington's. The idea is that by slippage of DNA polymerase CAG repeat areas can become so large that the transcribed protein cannot function properly and cause problems when it needs to be degraded by the proteosome. Eventually these defective proteins form aggragates that sequester all kinds of essential proteins, depleting the cell of functional proteins. Its just a matter of time before the cell can no longer cope with its disfunctional content.
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Recently, for my protein formulation stability experiment, I found massive visible particles in the high-temperature experiment. And massive particles detected  in micro-flow imaging(MFI) testing you can see attached file.Does anyone knows these particles? What are they?
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Tapan K Das, thanks.
And it's interesting that had not detected particles in the buffer solution(without containing protein).
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I am using ThT and Congo red assays to monitor Amyloid Beta 1-42 aggregation. It worked fine before, but now I am evaluating compounds with overlapping UV absorabance to these dyes. So any suggestions to alternative assays ?
Thanks
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Here is a link to a paper where we tested a range of different fluorescent dyes ThT and Congo Red against amyloids. Hope it is what you were looking for.
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This is a question my lab mate is working through:
Hi, I have an issue with RNA EMSA. I see the decrease in the intensity of the free probe with increase in protein concentration, but I barely see shift. There is also no increase in the intensity of the shifted band with the increase in protein concentration.
Here are conditions of my EMSA: For binding assay I use Binding buffer (10 mM Tris pH= 7.5, 10mM MgCl2, 100 mM KCl, 7.5% glycerol, 20 mM DTT), 250pM of probe (45), 4U RNase inhibitor, 2 ug tRNA and various concentrations of CsrA protein. Incubate for 30 min at 37C, then  run on 6% non-denaturing polyacrylamide gel in 0.5X TBE (pH~8.3-8.5) at 4C for 1h. Predicted pI of the protein is 8.16-8.3. Size of the protein monomer is 7kDa  and 14kDa of dimer. I used DLS to check if protein aggregates in the binding buffer, and it is not the case.
Attached is the typical picture I get. Why do you think the band is faint and seems to be stuck in the well? Thank you!
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Hi,
This is the issue that I have faced as well. I read some papers which suggest to increase the pH above the Ip (Isoelectric point) of the protein which gives it negative charge. However, I also found an article which suggests to decrease pH to obtain sharp bands.
@Peter Rehbein, I am wondering, if the pH is lower than the protein's Ip, meaning that it will have positive charge, how it can move through the gel from negative to positive?
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hi to all,
i have a stock solution of an ab amyloid peptide previously treated according to the standard protocols. this solution has been stocked at -20°C for a couple of years, and the peptide is no more in a monomeric state and is not properly folded. i was wandering if it is possible to restore the monomeric state and the correct folding with such treatment. unfortunately all the protocols i found are for the liophylized peptide, and not for a peptide solution.
thank you
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The protein by this time is almost certainly in the aggregated state. You could try to recover it by:
  1. Lyophilizing the sample to remove the solvent. Lyophillization is really the only choice here since your aggregated protein will clog any potential columns you could use.
  2. Redissolve the sample in 1% NH4OH at 4C to a concentration of less than 1mg/ml to disaggregate the sample. You can monitor the disaggregation process on Nanodrop. If the sample is monomeric, the Tyr peak at 276 nm should be clearly detectable as a distinct peak. If it is aggregated, the Tyr peak will obscured by the Rayleigh scattering from the aggregates. 
  3. Relyophillize the sample once finished. 
  4. Run the sample through a reverse phase HPLC
  5. Repeat steps 1-3 (the organic phase in reverse phase HPLC can trigger aggregation).
The reverse phase HPLC step is important at this point since you could no longer assume the protein is chemically pure. Met35 is likely oxidized and it is possible the peptide has undergone other modifications during the long storage period, such as deamidation, which can have a strong impact on aggregation kinetics and structures. [1,2]
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During protein concentration step by using Amicon tubes with 10KDa cutoff (from Merck company), I could see some cloudy suspension which indicate for protein aggregates formation. Afterwards in SEC (GF) step I had an elution profile with low resolution due to these aggregates which came in the void peak and start smearing as a shoulder for the void peak in the chromatogram.
I use DTT (1mM), EGTA (1mM), NaCl (50mM), and MgCl2 (1mM) in Lysis, wash , and gel filtration buffers
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Although very common due to its simplicity, concentration with centrifuge devices is possible the worst method available due to the extremely high local concentration you obtain at the membrane: many proteins aggregate or denature at such high concentrations. More delicate concentration methods are stirred devices (e.g. Amicon) or Tangential Flow Filtration or TFF (e.g. Sartorius).
Additionally, never inject dirty material on a chromatographic column, you should filter before the actual purification step, with 2 advantages: 1. remove the aggregated protein, and at the end of SEC obtain cleaner material; 2. prevent damage to your SEC column.
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A recent discussion amongst colleagues brought this question up and I hope some of you can help me here. We have been taught for a long time that Arginine as an amino acid is positively charged. A recent discussion ( mostly heated) involved role of an arginine amino in an active site of an enzyme. Most of the people agree that arginine is positively charged but disagreed on its role in electrostatic and hydrogen bond interactions. Because of the conjugation between the double bond and the nitrogen lone pairs in the side chain, the positive charge is delocalized.This means that the electrons are not associated to one atom but are free to move around...Now here lies the confusion.. Is it a golden rule that delocalized electrons cannot participate in eletrostatic interactions? How does this rule affect arginine? If the positive charge cannot play its role as a "charge" then how does the molecular interaction work?
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Hi there,
The case for the guanidinium of arginine is similar to the case of carboxylate in acidic aa (possibility of charge delocalization). They are both able to participate to salt bridge interaction which is actually made of both electrostatic interaction and hydrogen bonding contributions. Indeed global charge of arginine is delocalized between the 3 nitrogens of the guanidinium group but electrostatic interaction  may actually contribute to stabilize one of its resonance forms.
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enzyme is a terpenoid cyclase.
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I gather that the 1% Tween is needed to prevent the protein from aggregating. If the SEC column was not equilibrated with Tween, the concentration of Tween that eluted with the protein may have been too low to prevent aggregation. To make sure the Tween concentration remains at 1%, put 1% Tween in the buffer used to equilibrate the column.
You may not need such a high concentration of Tween, however. It may be sufficient to use 0.1% or even less once the protein is largely purified, as long as it isn't too concentrated.
If the protein is incorporated into the Tween micelles, it will elute at a larger apparent molecular weight because of the size of the micelles. You may conclude that the protein is aggregated even if it isn't.
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Hi all, I am measuring UV absorption spectra of whey protein by a UV/Vis spectrophotometer. Sample is used as 10 g/L without any dilutions, that gives me a range of absorbance from 1-2.5. Is this range of absorbance is acceptable? Thanks in advance. 
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Hi there,
As a rough estimate, I would expect an OD of 10 at 280nm for an unknown average protein solution at 10g/L.  But an OD of 10 can't be read directly on a classical spec so dilution is needed.
Other things to think of : have you run the Baseline first? What are the OD at 340nm (if not zero there is aggregation) and 260nm (ratio 260/280 should be around 0.6 if there is only protein in your sample)? What is the shape of the Spectrum? is it flatten at 280nm? If so the signal is saturating (so dilution is required prior Spectrum). Is your protein pure and do you know its sequence? If yes you can calculate its specific response factor at 280nm.
So you read 1 to 2.5 for 10g/L. This is low unless your protein is very poor in Tyr, Trp, Cys and Phe (these are aa contributing to OD at 280nm. If there is aggregation you should observe significant absorbance between 300 and 340nm.
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To isolate protein aggregates from neuronal cultures
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Some RIPA buffer recipes already contain SDS and deoxycholate detergents, both of which are anionic.
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I have doing ThT fluorescence (one point and time kinetics) on abeta peptides. My peptide is abeta 25-35 fragment. I am dissolving 1mg of it in 50ul 100% DMSO and then adding 225ul h20 and 225 ul DMEM buffer. final volume of this mixture is 500ul. I store this solution in 4 degrees. after 24 hours, I take it out.
I take out 110ul of this samples add 10ul (100uM ThT) and add 120 ul of this mix into a well of 384 well black plate (Nuanc) to take one point ThT fluorescence reading. I have also done experiments with other buffers and conditions. the observation I make with DMEM is that it has high fluorescence at 500-550. I compared this data with other conditions and did not find such observation.
Please kindly help me out what is going with abeta 25-35 fragment in DMEM compared to other buffer.
why I am using DMEM?
I am using DMEM or F12 can also be used as I read in papers that it is used specifically to produce oligomers of abeta 42 full length. People have used DMEM or F12 added into 5mM DMSO dissolved abeta 42 and kept at 4 degrees for 24 hours and observe oligomers by AFM.
Since abeta 25 is equivalent in aggregation and cytotoxicity to abeta 42 full lenth that is why I am using the same conditions to observe oligomers of abeta 25.
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Perhaps the other groups have normalized their data to take out the contribution of DMEM fluorescence? If the fluorescence you measured for DMEM alone (150 AU) is subtracted from the Abeta-DMEM measurement in the attached excel data (green curve) then the value is close to the other measurements.  I also noticed a red-shift in the Abeta-DMEM peak. Maybe there is something else in the DMEM contributing to fluorescence. There are many types of DMEM available.
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Can I use Dynamic Light Scattering Technique to determine protein aggregation?
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DLS is a quick way to measure protein aggregation without using much sample, and the sample can be recovered. It is very sensitive for detecting aggregation of protein. Software allows you to calculate the mass distribution, so you can get an idea of how much of the protein is aggregated.
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I have a 96 well plate set up and I am working with U87 cells. I would like to stain and image because I want to see the protein aggregates that form due to a compound that the cells are treated with.
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Since one is lost a bit in uncertainty about the "protein aggregates" expected to be stained (do you have ANY Thioflavin - Thioflavine* T or S ) staining protocol already? if yes would you mind to tell us?)  
as short as possible:  - try first:
and - perhaps for ending up with a decision for at least a first staining experiment:: stainsfile.info/StainsFile/stain/amyloid/fluorescent.htm
If you further are in doubt, try https://scholar.google.com searching for
< Difference between Thioflavin Thioflavine S and thioflavine T >
< Difference between Thioflavin Thioflavine S and thioflavine T staining > or
< Difference between Thioflavin S & thioflavin T staining > (different search phrases resulting in different result-numbers)
another source with general information on Thioflavin T:
"Thioflavin S: also referred to as Direct Yellow or primuline (and may be spelled thioflavin/thioflavine), "     = Interesting blog reading at:
Best wishes and good luck!
NB: *) the dye-name at least in German is: "THIOFLAVIN" (search for "thioflavin" in Scholar.Google yields 23.900 results, but many articles (titles) contain the name <thioflavine> (search in Scholar.Google yields 7.250 results).  Interestingly the information presented in Wikipedia (https://en.wikipedia.org/wiki/Thioflavin) does not contain the name "thioflavine"....but describes properties of Thioflavin T as well as Thioflavin S  {for the latter:  citation  "However unlike thioflavin T, it does not produce a characteristic shift in the excitation or emission spectra.[Lit.] This latter characteristic of thioflavin S results in high background fluorescence, making it unable to be used in quantitative measurements of fibril solutions.[Lit.]" end of citation} 
(The Wikipedia-page last modified on 15 July 2016, at 22:49).
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Hi, I want to show a colocalization between misfolded protein and endoplasmic reticulum but I'm not sure about the ER marker I should use for that ? I first wanted to use calreticulin or calnexin but i'm afraid that since they are chaperonnes their expression will change between my control and my "misfolded protein" condition. Does anybody have an advice ?
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Hi there,
What about anti-KDEL or anti-PDI antibodies?
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My protein of interest gets detected in IF, but not by western. Have used different tags. GFP and HA. Both works fine in IF. Protein is expressed and localized properly. No cell death or protein aggregation/degradation issues, based on IF info. Absolutely no signal in untransfected cells. Transfection efficiency was fairly good- 50-60%.
But only the GFP fusion gets detected in western, but not the HA-tag fusion. Absolutely blank lanes, no sign of degradation products/smear either. The same insert got cloned into both vectors. Both were N' fusions and were sequenced to confirm their fusion and frame. No question of epitope masking issue as it is an SDS-PAGE-Western.
IF on fixed tissues are a general problem with many antibodies that are good in westerns. An Ab that is good in IF, mostly works fine with open epitopes found in westerns. But this reverse and unusual problem we have is maddening and will be glad if anyone has any thoughts/suggestions.
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If your IF antibody recognizes a conformational or discontinuous epitope, then it would not be surprising that it doesn't work in a Western blot format. Do you have any detailed information about the epitope that's recognized by this antibody? If it is a commercial antibody the company should have some information about it.
Alternatively, your protein might adopt some non-natural conformation on the Western blot membrane, which is rare but not unheard off. You could try to add another denaturation step: after the transfer treat the membrane for 5 min in 1 N NaOH, then wash, block, and treat as usual. It might remove any non-natural conformation and render the epitope accessible. It has worked for me more than once.
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I'm trying to isolate and identify a protein from a fungal secretion based on it's activity.  I have tried GF using Superdex 75, Superdex 200 and Superose 6 columns but the only peak with activity comes in the void volume of the elution.  I've typically been using an elution buffer of 50 mM Tris, 100 mM NaCl (pH 8.5).  
I did get a glimpse of activity after the void volume one time when I tried upping the NaCl concentration to 500 mM, but wondering if I should try going to 1 M?  Also, should I try adding a detergent like Tween-20 or Triton X-100 to reduced aggregated protein?
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It's possible that the protein you are investigating normally exists as part of a high molecular weight complex or oligomer, rather than being aggregated. High salt treatment may disrupt this complex and allow the protein to be isolated by itself, but beware of the chaotropic (denaturing) effect of high NaCl concentrations. You may want to try a variety of salt concentrations to find the minimal effective concentration for eluting your protein after the void volume.
If you use detergent, you have to consider that proteins incorporated into micelles will elute at the molecular weight of the protein + micelle. In other words, the proteins will appear larger than they are. Some detergents have very small micelles (example: CHAPS) that minimize this problem. Also, make sure to use enough detergent that each protein molecule can be in its own micelle. Otherwise, proteins may coelute because they share a micelle rather than because they form a complex with each other.
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I am trying to purify an endonuclease (N-terminal hexahistidine tagged) of about 16kDa. The first round of purification involves overnight binding with Ni-NTA beads. When the elution fractions are pooled and injected for size exclusion chromatography, there is always a problem. 
1. Initially, my protein used to elute out in the void, that could possibly suggest aggregation, or an oligomeric state of high molecular weight. The yield was also very low. 
2. I had eventually begun adding NaCl to the buffers during batch purification.This slightly improved the yield, but after a gel filtration round, I see no protein at all. (Tried with both Superose 12 and Superdex 200 columns). 
This has been totally baffling me as the columns are working perfectly fine and yet I can't find the reason for my protein loss. My questions are:
1. What could be the reason for this loss of protein? I have read somewhere that sometimes the protein aggregates too much and non-specific adsorption may take place, resulting in protein loss. How do I eliminate that possibility? Can anybody tell me more about this problem?
2. And, if size exclusion is to be completely eliminated for purifying the protein, then how else can I purify the protein? Ion exchange is not working either. Is there anything else I can try?
Thank you so much!
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Hi Sravya
Size exclusion is not generally a method of choice early in a purification method. To be preparative, you need large columns and resolution is easily compromised. Gel filtration as a means to desalt or change buffer on the other hand is extremely useful throughout purification.
A word of general caution. New column media can produce very low yields during the first few runs. There is a certain amount of non-specific binding. Your mileage may very. 
As you are aware, the elution is related to shape, which if globular, is related to size. Aggregation is not uncommon and in some cases can be modulated with buffer conditions. I assume you are quantifying your protein with an accurate extinction coefficient and are calculating yields correctly. Are you monitoring fractions by hand or using a flow through monitor? Does the UV system measure at the wavelength your protein absorbs? I have seen many different kinds of mistakes made here. Just asking...
If there is aggregation, there is the possibility that the protein is stuck on the column. Do you see reduced flow properties or higher back pressure? 
As you have a his-tag, you should have 80-90+% pure protein - if you follow the tag and column instructions. How do you assess purity?
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I am doing tht test for assaying beta amyloid aggregation. I had two sample, one of them was "amyloid+ tht" as a positive control and the another was "my extract cell + amyloid+tht" and then we use 440 and 480 as excitaion and emission wavelenght.
But I saw that measured aggregation of beta amyloid at present cell extract is higher than the absent of cell extract.
In the other word, I want to know
1- why cell extract increases the beta amyloid aggregation?
2- Is it logical?
Please help me.
Thanks
 
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i use a lambda ex of 450 nm  and lambda emission of 482 nm 
i think you should make a control with your cell extract + ThT ..
and a simple advice.. you can filter the solution of tht before using it
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I am trying to concentrate the secretome of E. coli using Vivaspin columns (GE). I succesfully concentrate the proteins but when running SDS-PAGE I observe pretty blurry bands and irregular migration, presumably result of protein aggregation while concentration. I have tried to shorten centrifugation time and include sample mixing between centrifugation rounds, but gels do not look much better. The buffer where I am trying to concentrate the proteins (LB medium itself) is definitely not ideal, but:
1: Could you provide me with tips in terms of temperature, additives ... to reduce protein aggregation and improve the quality of the concentrated protein sample?
2: Could I somehow reverse the aggregates in my samples?
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 First assume that you are running protein MW standards, and they run fine on the same gel that your samples do not "and" the MW standards have the same SDS/PAGE sample buffer added to them as do your protein samples.
1) If you are running the gels just for analysis purposes, then make the samples 8 M urea "before" adding the SDS/PAGE sample buffer and if you heat them do not go above 95oC.
2) For proteins that have the tendency to aggregate during concentration, try using the Millipore tangential flow filtration (TFF)-based filters, they reduce the generation of a steep concentration gradient during concentration and so reduce aggregate formation, the alternative is to use a tangential flow filtration (TFF)-based concentration cell, but then you need a N2 tank with regulator, though the cell will eliminate any concentration gradient from forming.
3) Have you treated the samples with a nuclease?, and analyzed the protein sample for DNA/RNA? (A260/A280 ratio). DNA can contribute to aggregation of proteins at all steps of purification and concentration.
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I am doing tht test for assaying beta amyloid aggregation. I had two sample, one of them was "amyloid+ tht" as a positive control and the another was "my extract cell + amyloid+tht" and then we use 440 and 480 as excitaion and emission wavelenght.
But I saw that measured aggregation of beta amyloid at present cell extract is higher than the absent of cell extract.
In the other word, I want to know
1- why cell extract increases the beta amyloid aggregation?
2- Is it logical?
Please help me.
Thanks
 
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First of all you should make a control consisting of cell extract plus Tht. Cell extract is a very complex mixture of molecules that can bind Tht and change is emission spectrum. This "blank" should be subtracted from your experimental sample.
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Hi all,
I am working on designing insoluble proteins and their behaviors.
What I want to know is "Can I use the dynamic light scattering (DLS) to see the aggregation properties even if my proteins are not soluble in the buffers?"
What I actually did was I modified the genes in crystalline forming region of my proteins, so I do believe the their aggregation should be changed when I express the proteins with e. coli.
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You can probably learn something about the aggregation of your modified protein, but it may be quite tricky. If the protein is entirely insoluble then it would not be possible to do DLS unless it is dispersed somehow (for example with the help of surfactants). In reality, you may find that depending on your modification you observe different degrees of solubility, and DLS may help discern the level of solubility (average size and average polydispersity), for example versus temperature or when looking at different buffers/additives.
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I want to detect kinetic of beta amyloid aggregation in present of lysed cell of caco2 cell line and in present of beta amyloid? please help me
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Hi,
since you posted a question about advise on how to design an experiment it would be helpful to understand a bit better the broader questions you are trying to answer, but I`ll do my best with what I have. My guess is that you suspect something in this lysate will alter the aggregation process and/or kinetics. I have done similar experiments and they were straightforward. You mix Abeta with ThT and the substance which is expected to modify the aggregation and compare the time dependent ThT signal to a control with only Abeta and ThT. You can record the fluorescence in real time in a cuvette (also you might have some sedimentation effects of larger aggregates that will make your solution inhomogeneous, which can be prevented by stirring the solution, but this leads to other problems like sheer forces on the aggregates, creating air bubbles that scatter light and similar things that aren´t necessarily reproducable) or you can take samples in regular time intervals of your aggregating sample (after mixing it well) to reduce the sedimentation effects.
Where it gets messy is if you add the whole lysate. So if you have any idea what it is about the lysate that possibly modifies the aggregation kinetics, I would recommend to isolate it so that you will have a better defined system. If you add the whole lysate there are so many players that can interfere with the aggregation and the ThT-fluorescence and of course also the physical properties of the solution would change (the pH you can easily adjust in your control sample, but also the thermodynamic activity of water would change and you would have a vast number of interaction surfaces your key players are exposed to). If you can´t isolate your key component in the lysate or at least fractionate it, my two major concerns you should at least check for would be:
1. ThT can in my experience interact with many things. Everything that restricts the rotation of the two ringsystems increases its quantum yield. I guess if your starting fluorescence (of course with the same dye concentration) is the same you can assume that ThT isn´t bound to anything randomly present in your lysate (also you can´t be 100% certain.
2. The Abeta aggregation might already be altered just by the presence of other substances in solution. Higher crowding and availability of hydrophobic surfaces might push the protein to the aggregated state and therefore enhance the kinetic. This is difficult to correct for, so if you want to check for a specific effect of one interaction it is hard to separate these effects.
2.
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Supernatant proceed to ammonium sulphate precipitation (standard method)
1)      0-20% (centrifuge 7000g x 20 min) -> 20%-50% (centrifuge 7000g x 20 min)
2)      Procedure done in ice while stirring
3)      Fine ammonium sulphate                 
4)      supernatant was buffered with 50mM Tris-HCl pH7.5 prior experiment
5)      (NH4)2SO4 was added slowly (17.55 g in 50ml ), fully added between 5-8hrs (white foam cannot be avoid)
6)      Incubated at 4 ͦC chiller for 12hrs while stirring
Note: activity loss  about 75%, fold less than 1
Q: do i need to add all the ammonium sulphate within certain range of time and how can i avoid formation of white foam (very little)?
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You might consider HIC chromatography in lieu of precipitation.  But is the basic question concerning protein folding? If it's about folding:   It may not matter how you purify the protein if you intend to refold as the next stage.  Refolding usually begins with complete protein denaturation firstly followed by a partial factorial matrix to determine folding conditions.  So if your purification results in denaturation, you would be denaturing the protein anyway to start refolds. 
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Dear all,
my E. coli lysate is extremely viscous, to the point that it is impossible to apply it to the column for purification. What can I do?
I added 500 units of DNAse to 50 ml, but the resulting suspension was almost as viscous as honey... :/
I am restricted to chemical lysis with detergent, for safety reasons (potentially infectious protein). So no french press for shearing DNA is applicable...
I read that working on ice throughout the lysis process would allow for spinning down the DNA in the centrifuge, is this true?
Any help on this is highly appreciated. Thanks in advance for any advice!
Best,
Peter
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To reduce viscosity due to RNA and DNA I typically use Benzonase from Sigma (http://www.sigmaaldrich.com/catalog/product/sigma/e1014?lang=en&region=US). It is far more effective on DNA and RNA and works well at pH 7.0-8.0.  It is superior to DNase for the procedure that you are performing. In addition, you could always treat with Benzonase and then run your sample through a syringe with a small gauge needle on the end to sheer any existing polynucleotides to further reduce the viscosity. 
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I'm looking for alpha synuclein aggregates.   I have a few questions about the technique.
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