Mass Spectrometry - Science method

In my case, I guess I have used any one 5-carbon compound and 6-carbon compound. I guess you can switch among them based on your interest and instrument specifications.

Please go through the Internal Standards:

How about protein A or G resins...The melon gel works as flow-through mode of purification, protein A is bind & elute mode (orthogonal strategies). Without knowing about a cost comparison, this application works well and recovery would be satisfying. If you investigate the IgG CDR domains (fab) nsmol kit is ready to use and a promising tool as a front-end approach.

Hi Himani,

If you know which organisms are included in your samples you can work with a joined database fused from downloaded fasta format databases (i.e. from Uniprot) of the respective organisms.

With Maxquant you can than look at the number of unique/razor peptides to check from which organisms the proteins are more likely.

Best,

Murat

Hi Dina,

What is the purpose of the analysis your performing? If you'd like to get an idea of the yield of a protein purification, I don't think you would need a protein digestion protocol; you could consider running LC-UV instead.

How did you determine the yield of the individual steps of your protocol?

And how are you desalting after digesting?

What is your 'running solution'?

Thanks in advance for providing some additional context and information!

As an update to this discussion, I have decided to reduce my sample size and incorporate a pooled reference channel. Mostly to open up the possibility of integrating additional samples and conditions in the future.

Sam

When performing mass spectrometry analysis of lipid classes using a lipidomics standard like the EQUISPLASH product from Avanti Polar Lipids, adding the standard to your test sample serves as an internal standard for quantification. This approach helps account for variations that can occur during sample preparation, extraction, and analysis. Here's how it works:

1. Internal Standard Principle:An internal standard is a compound that is added to both your standard samples and your test samples in a known amount. It's chemically similar to the analytes of interest but should be easily distinguishable in the mass spectrometry analysis. By adding a known amount of the internal standard, you can correct for variations that might occur during sample preparation and analysis.

2. Adding the Internal Standard:When you add the EQUISPLASH lipidomics standard to your test sample during the extraction process, it becomes a reference compound with a known concentration that you control. This internal standard will undergo the same extraction and analysis steps as your sample, which helps compensate for any losses, variations, or biases introduced during these steps.

3. Quantitative Data Analysis:To obtain quantitative data from your mass spectrometry analysis, you'll follow these steps:

The rationale behind this is that the internal standard's known concentration serves as a reference point. If the extraction and analysis are consistent, the ratio of the analyte's peak area to the internal standard's peak area should be proportional to the ratio of their concentrations.

4. Calibration Curve:To convert the peak area ratio to quantitative concentration, you'll need to create a calibration curve. This involves analyzing a series of standard solutions with known concentrations of the lipid classes using the same extraction and analysis procedures. The calibration curve plots the concentration of the internal standard against the measured peak area ratio.

5. Quantification:Using the calibration curve, you can then determine the concentration of your lipid classes in your test samples based on the peak area ratio. The formula to calculate the concentration is derived from the linear relationship shown in the calibration curve.

By adding the internal standard and utilizing a calibration curve, you can obtain quantitative data for your lipid classes in your test samples, even considering any variations introduced during sample preparation and analysis. This internal standard approach enhances the accuracy and reliability of your quantitative results. Subhadip Kundu

To get qualified and specific answers you need to provide a bit more information about your experimental approach. Ismail suggested many different options that may be suitable depending on what your overall experimental setup is. As i read your question you are capturing interactors on a Ni-NTA resin with the bait protein bound and then eluting them with imidazole.

There are a wealth of proteomics sample prep methods nowadays that would get you past the imidazole issues, such as protein aggregation capture (PAC) or SP3, which serve to precipitate your protein onto a bead matrix that can be washed thoroughly before trypsin digestion (if bottom-up proteomics is your method). Alternatively, you may be able to digest your protein directly on the Ni-NTA matrix. A C18 desalting step is usually standard procedure before LC-MS analysis and that should give you nice clean samples overall. Gel-band analysis is another approach that should also work fine (if a bit more laborious).

I would suggest your consult with a proteomics specialist at your institution to figure out the most viable approach for your experiment.

The best way for this is to use the MALDI-TOF MS configuration. For ESI top-down or native-MS-like approach is needed for the analysis of either heavy or light chains. You may combine SEC and native analysis for ESI-LCMS or gas phase fragmentation of antibody light chain to select SRM-like produced peptide for quantification. ESI produces multiple charge states for large molecules therefore occurring charge envelopes reduce the precursor intensity if the top-down quantification is aimed. MALDI gives reduced charge states thus either the identification or quantification approach would be more easy/more practical if the sample is not a protein complex but a purified antibody.

Rather than the selection of MS or acquisition technique, it is more important to choose the sample prep strategy herein. Garbage in garbage out for any MS system. What is your consideration about the sample prep and what is your sample matrix? how would you purify/clean, reduce, and fractionate your sample? It is more critical to assess prior to MS detection. Otherwise many interfering compounds, and protein peptides make your quantification worse and that is why it is recommended to use MRM and signature proteolytic peptide identification is more appropriate to perform absolute quantification...

Alternatively go back over your sample preparation protocol and determine the source of the PEG contamination.

If your laboratory results are showing increased zinc levels despite collecting samples in EDTA tubes, there could be several potential reasons for this discrepancy:

To resolve the discrepancy and ensure accurate results, it is essential to review the entire testing process, including sample collection, handling, and analytical methodology. If there are concerns about the results, it is recommended to consult with the laboratory or a qualified healthcare professional for further investigation and appropriate interpretation of the findings.

Thanks Murat

I suggest using phree-phenomenex (bimodal/phospholipids and protein removal) protein precipitation plates...This will remove the abundant large molecules of serum such as proteins (albumins, globulins...and phospholipid derivates)...with this, you may combine protein precipitation and spe clean-up into a single prep step...In addition, MS/MS fragmentation and using alternative ions for your SRM acquisition may improve specificity and give more non-interfered spectra...If sensitivity is low I also recommend nitrogen evoporation followed by buffer exchange and internal standard normalization approach for absolute quantification...for hydrophobic compounds such as curcumin I also use APCI instead of ESI to get a more intense signal if your configuration is applicable to implement...

Good luck

No. You can never assume that different compounds will ionize in the same manner. MS analysis is not a "universal" detection technique.

Hi, I don't know the extent of your shiny application, but you can also use EnrichR (in R), which is also good, though, it upload the data to server during calculation.

Plus, I am curious how you will handle the missing IDs during ID mapping? ClusterProfiler can map IDs, but there are always some % of ID that are missing.

I would extend that and say we really need to be careful using upregulated. To me, that means the expression of the protein is increased, so there is a fundamental change in the amount of protein quantified as a result of that. I'd suggest we use the term increased (or decreased) abundance when quantifying the protein, unless we have clear evidence to say otherwise. There are a lot of reasons protein quant differs between samples, and it's not always due to expression level changes. I agree you will find such terms used interchangeably, that does not mean it's correct, or a good idea.

Yes, this is flying as a protonated adduct.

Noel W Davies thank you very much

Dear Hongjie Hu,

Please share the data it will easy to discuss.

I would do an on-bead digestion with trypsin, which will give you the peptides that can then be followed by cleanup (if needed) and MS. This is quite a common approach for elution and any mass spec service should be able to advise you on it.

To my understanding, acetonitrile, methanol, isopropoanol etc. act as electron donors in ESI conditions, enhancing ionization in ESI+. This is one of the reason why ionization in ESI+ is enhanced by solvents (the other is their faster vaporization than water in ESI conditions).

some solvents are better electron donors than others, and you may notice signal enhancement switching from one to another...

hope this helps...

Dear Simon Thain,

Thank you for the answer! Yes, our case was caused by losing BIOS settings, we restored them and now Synapt is running. But absence of the manual for EPC is looks strange for me. battery in EPC is not eternal, so why did they don't add any information in user manual or even their web-site? It looks like Waters don't even think about situation, when their instrument will be online after more than 5 years :-)

Warm regards,

Azamat

Noel W Davies

Thank you so much Prof. Noel

Let me work on your advice and will come back to you for feedback.

Thanks

İsmail Emir Akyildiz that is really a good suggestion. I would try this out

Dynamic light scattering (DLS) and small-angle X-ray scattering (SAXS) are pretty much universal techniques. AFM might be demanding for sample preparation, but it is also possible.

Shift in the surface plasmon resonance (SFR) UV-VIS absorption peak works in some specific cases.

Dear Itay,

do you see no signals at all, or is the identification/quantification of sugars connected to peptides the problem (e.g. in a software) ? Do you have a chromatography in front or do you only want to analyse single proteins and thus have a direct injection setup?

Yes, there are several other metabolome databases available aside from the Human Metabolome Database (HMDB). Here are some examples:

Metlin: A metabolite database that includes both experimental and predicted mass spectrometry data for a wide range of metabolites, including those from human and other species.

KEGG: A comprehensive resource for understanding metabolic pathways in various organisms, including human. It contains information on biochemical pathways, enzymes, and metabolites.

LipidMaps: A database of lipids, including information on their structures, physical properties, and associated enzymes and pathways.

BioCyc: A collection of databases that includes information on metabolic pathways, enzymes, and metabolites for a range of organisms.

ECMDB: The E. coli Metabolome Database, which provides information on the metabolites and metabolic pathways of Escherichia coli.

MetaboLights: A database of metabolomics studies, containing information on experimental designs, protocols, and raw data, as well as curated metadata and metabolite annotations.

These are just a few examples of the many metabolome databases that are available. Each database has its own strengths and limitations, and the choice of database will depend on the specific research question or application.

There are several causes for the presence of outlyers. Impurities, errors during the extraction, the needle that for some reason did not inject the same amount of sample, and samples that are naturally different from the other ones.

You can solve the problem using this checklist.

- check the look of the chromatograms and verify that all of them look more or less the same.

- check IS intensities and verify that their peak area is within an acceptable range for all the samples.

- normalize the data across samples.

- scale the data using UV scaling.

- evaluate the presence of outlyers using RobustPCA.

good luck

Late answer but TiO2 and FeNTA are not suitable for phosphotyrosines( i.e I quantified 43 phosphotyrosines by a sample from which I quantified nearly 9000 S/T phosphosites so much less than 1% with TiO2) If you don't have sample quantity problems(not working on clinical data) pY100 antibodies are the best for phosphotyrosines. Not really an expert but if you can quantify more than 300 phosphotyrosines its a success.

You need to freeze them immediately using liquid nitrogen in order to stop the metabolism of the cell. After this you can keep the cells until the analysis day.

routing problem, contact service engineers of Thermofisher.

Dear Khawla,

former colleagues of mine created this database for biomarkers: http://bionda.mpc.ruhr-uni-bochum.de/start.php

You can either look for a disease and find biomarkers or directly for a protein of your choice to see if it was identified as being a biomarker previously.

However as Jennifer pointed out especially RT values are highly dependent on the chromatography was used and can differ significantly. The m/z value should be identical for the majority of the MS machines, can differ though, when differently charged ion species are prefered by MS method or application. Nowadays there are also a lot of predicition tools in proteomics which help you to predict this values for different set ups. This might also be an option.

Given only that information it is very difficult to make a tentative identification of the compound, you have to explain what specific analysis was performed if it was only masses, or masses masses and there are fragments that can further support the identification. Also, information about the compounds previously identified in that plant could help you to know what type of compound it is. I leave you some pages that may help you to identify the compounds.

Hi Yang,

Is the sequence of the protein you're interested in known? In other words. would you like to confirm its sequence, or do you mean you want to de novo sequence it?

My first response would to proteolytically digest it and analyze the resulting mixture of peptides with LC-MS/MS to get accurate masses and fragmentation. In this way, you should be able to reconstruct the amino acid sequence. Getting full sequence coverage sometimes requires the use of multiple proteolytic enzymes.

Another possibility would be to top-down sequence it, for example using electron transfer dissociation, but in that case you'd need access to that type of equipment.

So, basically, a few remaining questions are:

- what do you know of your protein of interest already (also regarding modifications, such as glycosylation)?

- what type of MS equipment do you have available?

Mrs/Miss Saeedi,

There is needed only raw of your measurable variables.

There can be used software: [https://www.thermofisher.com/order/catalog/product/de/de/10137985].

Please, see our work on MALDI [1].

[1] Journal of Molecular Structure, 1260 (2022) 132701

Stochastic dynamic quantitative and 3D structural matrix assisted laser desorption/ionization mass spectrometric analyses of mixture of nucleosides

Bojidarka Ivanova, Michael Spiteller

There is no exact stoichiometry for any large cycle in metabolic networks. You can write out the system of equations for all the potential metabolites, then you need to add the recycle equations (including oxidative phosphorylation and PMF build up from NADH recycle). You will end up with more equations than unknowns, which implies an infinite number of solutions. To this system you need to add the sum of the free energies of all the reactions Delta Gr ≤ 0 to get the reactions to proceed, again, the inequality isn't sufficient to define a unique solution, only a boundary beyond which certain stoichiometries can't exist. This issue is called metabolic uncoupling. An example of this (for mixed acid fermentation), which explains the issues in biochemical stoichiometries, is provided in Schneider, LV, Biological Engineering: The unit operations and mathematical modeling of biology, Chp 2, (LVS Sciences, Houston, 2022). You can also look at the online lecture from this book (https://www.youtube.com/watch?v=6r9MVzwxdAs)

Mr. Siddique,

Exact quantitative analysis of any analytes by means of mass spectrometry can be carried out, currently, only within the framework of our (own authored theory to me and my co-author as shown in the corresponding references, below) innovative mass spectrometric strochastic dynamic theory and model equations.

Please, consider works [1,3] and the reference section, therein.

[1] Steroids, 181, 2022, 109001

Mass spectrometric stochastic dynamic 3D structural analysis of mixture of steroids in solution – Experimental and theoretical study

B. Ivanova, M. Spiteller

[2] B. Ivanova, M. Spiteller, Stochastic dynamic ultraviolet photofragmentation and high collision energy dissociation mass spectrometric kinetics of triadimenol and sucralose, 2022, in press.

[3] B. Ivanova, M. Spiteller, Exact quantifying of mass spectrometric variable intensity of analyte peaks with respect to experimental conditions of measurements – a stochastic dynamic approach, 2022, in press.

Particularly, an examination of peptides via our approach we have performed, as well. Please, consider work [4].

However, please, be informed that the latter contribution shows only application of the basic stochastic dynamic equation. The next two (2) derivative equations within the framework of the same theory and basic model formula are not used in this study.

[4] B. Ivanova, M. Spiteller, Chapter 1. Experimental Mass Spectrometric and Theoretical Treatment of the Effect of Protonation on the 3D Molecular and Electronic Structures of Low Molecular Weight Organics and Metal–Organics of Silver(I) Ion, In Protonation: Properties, Applications and Effects, A. Germogen (Ed.) NOVA Science Publishing, New York, 2019.

Firstly, in SALDI experiments, a protonated ion can hardly be observed. Once you found a [M+H]+, we would like to consider that the H source may come from the containment of your MALDI target, or it can come from the acid species of your sample.

Hi Sam,

your approach depends on how many samples? If it can fit into a single TMT run, and I’m pretty sure they are up to 29-plex now, then you’ll be hard pressed to beat variability with DIA. If you are running >60 samples (so now you have at least 3 x 29-plex runs) then I would argue DIA would be preferable. You clearly have the MS to do either approach. Also consider the expertise in the lab for both running the samples and analysing the data.

If you do a search, you’ll find plenty of papers comparing DIA, to TMT, to DDA. This tends not to be batch variability comparison, more coverage and missing values. There has also been a lot of work recently on removing batch affects and unless you have a shocker, it can be removed in most cases.

Good luck,

Peter

This method developed by Sili Fan and col. is a great option for batch correction.

Systematic Error Removal Using Random Forest for Normalizing Large-Scale Untargeted Lipidomics Data

DOI: 10.1021/acs.analchem.8b05592

Hi,

Can you explain what you mean by number of protein groups? Is that based on GO terms, for example?

There can be numerous reasons for why you find different data between different runs: did you analyze the exact same sample in both cases, or are these independent preparations of the same lysate? And how much time was in between the runs? Do you know how many peptides were detected in both runs, did you check what the TIC in both runs looks like, to get an idea of overall data quality comparability between runs? Have other people been running samples in the meantime?

Depending on the calibration of the MS, the overall cleanliness of the system, etc., the data obtained from a mass spec experiment can greatly differ.. A bit more information would be helpful, I guess.

Hi Adela,

I missed your response. Yes, I have measured the linearity of both the detector as well as the depletion of the ion beam with the power of the laser at several wavelengths. The issue is that day-to-day I cannot get consistent overlap of the ion/laser beams and the optical path is not well-aligned (old vacuum parts--I suspect one cross flange is bent by 1-2 degrees). The OPO laser beam is roughly 1x0.5 cm^2 and when scanning from UV to IR the most intense part of the beam walks around a bit. An iris aperture and a detector near the interaction region is my best idea of solving the issue.

Hello i really appreciate your help ! I tried before that transformation but it was not the same as compared in R studio. I thought maybe there is a formula that i can put manually to do that.

In mass spectrometry data, switching between data scales can change the whole interpretation of the results. For that reason i want to apply the same transformation as my team do with Rstudio..

There are also the T2 system and the BioFire FilmArray system.

Don't be surprised that plasma (the sample matrix) will interfere with HPLC-UV detection. So the answer depends on 'which' amino acids and their concentration.

Various instruments and methods have different requirements for sample volume.

In the case of classic autosamplers there is some minimal vial volume. If you choose the right vials with special inserts, the minimal sample volume would be about 10 ul.

There are some tricks to avoid this problem, e.g. elution of the sample from solid phase as in the Evosep system, but it is not very common yet.

In techniques such as MALDI ionization there is no minimal sample volume recruitments because the sample is mixing with a specialized matrix during sample preparation.

The question about sensitivity is much more complicated - different molecules will have various detection range in the same conditions. To maximize sensitivity you should carefully choose ionization conditions and try to avoid contaminations.

By our experience, in the case of MALDI, the purity of the sample often is much more important than amount of target peptide in it. Sometimes, when our colleagues use poor quality plastic and do not work properly it is impossible to obtain a good signal even in very concentrated sample. The quality of plastics and solvents is very important for mass spectrometry.

Other example comes from ESI-ionization. Addition of TFA instead of FA to the sample significantly reduces signal strength for most peptides.

Finally, do not forget about derivatization techniques. Derivatization is aimed to perform chemical modifications which significantly increase ionization efficiency of the target molecule.

https://www.sciencedirect.com/science/article/abs/pii/S016599361400096X#:~:text=Derivatization%20primarily%20aims%20to%20obtain,separation%20for%20some%20chiral%20drugs.

Thank you, Ismail and Eef. Finally I made it work using Protein Prospector. The drop-down list did not help as it only includes a limited number of modifications. What I did is to define new amino acids with isotope-labelling. For example, the isotope-labeled lysine can be defined as k = 13C6 H12 15N2 O1 (please note that one H2O need to be removed). Then the software can generate correct theoretical m/z for both the parent ion and fragment ions.

It is better to know what kind of modification you predict and how long the PTM peptide is. We may filter the troubleshoot after this information in terms of non-specific binding, or flow through elution. We can also suggest at this point enriching the PTM peptide by using an affinity-based technique such as TiO2 for phosphor proteome, lectin for glycoproteome, di-gly for ubiquitin, etc... This may be a simple dynamic range problem and can be fixed after a specific or nonspecific peptidome enrichment...

Good luck, emir...

Dear Xiaoyin Zeng,

MaxQuant is offering a Summer School every year, whose recordings you can also find on youtube. They have a complete tutorial on how to do PTM analysis of proteins. I am sure you will find your answers there.

Hi Sauradip Chaudhuri,

If you do an on beads digestion, you will only get peptides which will be cutted away from the Streptavidin bead/desthiobiotin complex. Depending on the digestion enzyme and the available digestion sites on desthiobiotin you may also have some desthiobiotin peptides in your identified sequences after LC-MS analysis but this should not be a problem if you perform your LC-MS analysis and downstream data analysis in the correct way.

Good luck!

Murat

Hello Yuhong Mao,

Perhaps try a few injections without the use of a trap column. This will identify if the analytical column is functional or not.

Once you have determined the analytical column is functional, add a trap column but install it so that the loading and eluting directions are the same i.e., do not operate in a reverse flush way as many columns no longer support this.

Finally, ensure that the column switching timings are correct so you have adequate time to transfer the sample to the analytical column.

Good luck.

Best,

Peter

The oldest of old school methods. By separating by 2D-PAGE, you need to get the protein out of the gel, which means either blotting to PVDF or ingel digestion to peptides. Blotting to PVDF means that your Edman will give you the amino acid sequence from the N-terminal which may be enough to give you a match. Ingel digestion means that you have peptides that you then need to separate by reversed phase and then Edman separately which you could then use to reassemble into the protein sequence. However, the won't be a difference in the results obtained if your sequencer can reliably give you 10-20 amino acids. But it will take 50-100x longer than LC-MS/MS. Edman cycles are typically 1 hour per amino acid whereas LC-MS/MS for a single spot is ~30 mins using nanoflow.

Adding to the complexity of this, there are actually many proteoforms in a single 2D separated spot.

Are you able to perform an in gel digest and send the peptides to someone who can do the LC-MS/MS for you? It would save an enormous amount of time and expense compared to the Edman.

It strictly depends on the overall approach to antibody sequencing. The main benefits of TIMSTOF instrument is high sensitivity and high ms/ms speed. It best fits to the transcriptome-assisted sequencing of native affinity purified pools( e.g. repertoire analysis of convalescent sera) It gives the acceptable quality ms2 of tryptic peptides with deep overall proteome coverage.

The same time timstof is inadequate instrument for true de-novo sequencing of mabs without database assistance. The modern Orbitraps performs multiple modes of fragmentation(CID, HCD, ETD, EThcD, UVPD) for different type proteolityc peptides. It greatly improves the quality of identification of long peptides (especially DMAPA-treated V8 peptides) and allows to differentiate Leu/Ile at the protein level. But this is true only for high-end Tribrid series Orbitraps(and partially for ETD hybrids) but not for Exactive and Exploris series.

Dear Jong Won Han,

our institute is based in Germany and we are faced with the same problem for several months now. We switched to steel needles, which are reusable as you can clean them in Methanole. The only drawback so far is that they are not recommended to be used for Phospho PTM analysis and additionally they are not compatible with a FAIMS device. Maybe steel needles could be an option for you to try, if your work is not focused on the drawbacks mentioned above.

Best wishes,

Britta

Try ISOGEOCHEM if you not already did.

https://list.uvm.edu/cgi-bin/wa?A0=isogeochem

As Tomas wrote "it's not uncommon to observe the interaction only in one direction"; all your prot A could be in the complexes whereas protein B and C are in vast excess in the cell and only a small fraction in complex with A, binding of antibodies to B could disrupt the interaction with A, or its epitope masked when in complex with A ....

Essentially, you have to open up the quads (both) and clean them periodically.

Dear Samit Karmakar,

THat was a part of the installation/device used in my very old work

The palladium window was used for dossage of tritium to the experimental volume by heating when T2 was to add.

Hiba Salim as I mentioned earlier, it is helpful to know what type of data you are receiving: is it deconvoluted mass of the molecular ion or fragment ions? If all you have from the measurement is molecular ion mass, then identifying this protein reliably will not be possible. There is, however, an option of checking for PTM from mass difference between predicted mass of the protein of interest and experimentally detected mass. Likewise, checking for protein degradation is possible by comparing detected mass against predicted fragmentation series. Good luck!

Depends on the standard chemistry and quantitation strategy. Can be chemical degradation due to storage, for example, or absorbance to vial walls. If abundance was assessed by MS signal, instrument performance specs on a tuning mix or something you use to benchmark performance need to be demonstrated before making conclusion on sample abundance.

Without background information I would presume that we will not able to help You... Sorry.

Hi Samantha,

Did you use Epoxy m270 dynabeads kit finally to avoid IgG contamination from IP samples?

If yes, how is your result?

Thanks!

As a very general rule of thumb, the protein yield after tissue extraction in SDS or urea buffers with sonication/bead beater for total cell lysis, including organelles (whole proteome extraction) will be approximately 5-10% of the initial wet weight. So from a piece of tissue that weighs 100 mg, one might expect 5-10 mg of protein. This varies of course depending on the type of tissue being processed, the stringency of the lysis/extraction reagents and process (are you pulverizing the frozen tissue first?), whether the tissue is dehydrated, and the volume of buffer used for lysis.

First of all, any IP will give you an enrichment, not a pure purification. So seeing many bands on an IP is not unusual. Even if you get a 100x enrichment, you will still have a lot of other bands in the lane besides your specific protein of interest.

For poorly soluble nuclear proteins you could prepare chromatin as you do for ChIP. Cross-link cells with formaldehyde then solubalize chromatin in 1% SDS buffer. This denatures all proteins, but if they are cross-linked then they will remain bound. Dilute the 1%SDS buffer about 1:10 for IP and then try with your antibody. It is essentially a ChIP assay but after IP and wash, analyze the proteins by reversing the cross-links and boiling in SDS/PAGE buffer. Check for your protein of interest and any bound co-IPed proteins by Western blot.

Comment on units: amu is an outdated unit, although it's still implemented in many softwares. The current unit is "u". It is:

1 amu = 1/16*m(¹⁶O)

1u = 1/12*m(¹²C)

The also often-used unit "Dalton" (Da) was originally defined as 1Da=1amu, but has been redefined as 1Da=1u. The fun part is that spectrometers that give out results in Dalton don't tell you whether the old or the new Dalton is implemented.

For most coarse measurements the difference between u and amu is within the instrumental accuracy, but if you're using an FTICR or an Orbitrap it makes a difference.

You can dissolve your analyte in (3:1) Chloroform/Methanol or (1:1)Methyl-tert butyl ether(MTBE)/water, these are mostly used solvents for lipid extraction for Mass spec analysis, you can keep the sample with solvent in a water sonication bath for couple of min & incubate @30°C for better result.

Dear Dr Naga Veera Yerra,

These softwares may actually help you (MetFrag and XCMS).

Here's their respective links:

Best wishes,

Sabri

Please see this file

Hi Ashish Bihani,

I hope all your queries get answered with these articles:

Thank you.

Varsha

شكرا جزيلا

You may add also lysozyme and deoxycholate as ms compatible surfactant along with phosphatase/protease inhibitors to lyse the cells. Deoxycholate is a membrane soluble anionic labile detergent and can be degraded in low pH conditions. Urea is also a good recommendation. However, you can perform the FASP protocol to get rid of MS incompatible detergents prior to analysis and simultaneously can digest and collect the phosphoprotein efficiently. Collected peptide fraction (filtrate) can be further used to selective enrichment of phosphopeptides by using commercial Phosphoprotein Enrichment Kit.

Dear Gabriela,

the Matrix Science website (Mascot database) has some good pdf files for training, which could be helpful for you.

Here is the link:

Good luck,

Murat