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No. Protein identification by mass spectrometry, unlike DNA sequencing, is a very complex process. Early discussion with the facility regarding the sample preparation process is necessary in order to obtain optimal results. Discussion of your experiment is required and is by appointment basis only.

During submission - provide as much information as possible about the sample, the background of the experiment, known facts, known and/or expected proteins, or any other information.

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As early as possible. Whenever you start planning the experiment, discuss with the facility about feasibility.

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First samples need to be registered by the user/student. An automatic email requesting approval will be sent to the PI after registration. Samples will be collected by the facility only after approval is received from the PI. Hence, approval from PI is MUST for sample submission. Please see "Sample Submission Guidelines" for detail.

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Our priority is to serve the internal users. However, collaborative use is supported. Internal users are requested to clearly indicate the source of the samples for collaborative work.

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Depends on the number of samples, complexity, LC gradient being used etc. Please discuss with the facility during sample submission to get a better answer.

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Please check the portal after sample submission at intervals. Portal is updated regularly.

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Depends very much on the protein!

Even a weakly Coomassie-stained band is enough for protein identification by nanoLC-MS/MS. But, proteins are different, the success of protein identification also depends on the protein itself (its amino acid composition, hydrophobicity etc), some proteins can be identified at very low level, some are MS incompatible.

Before you start your experiments, discuss with the facility and give as much information as possible.

Prior to MS analysis, as standard practice, we use an internal control of a tryptic BSA digest at a level of 100 fmole, which is almost the lowest level visible by colloidal Coomassie. This injection gives sequence coverage of over 50% and a high database search score. The limit of detection of the instruments for BSA is below 10 fmole.

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The total amount is between 20 - 50 µg/lane, but it also depends on the complexity of the sample. In general, good Coomassie-stained gels with clear bands are best for consecutive analysis.

Overload gels cause problems, because high abundant proteins will be seen in many bands and mask the "interesting" low abundant proteins.

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1) Polyethyleneglycol (PEG) and detergents containing PEG (common pegylated detergents include; Triton X-100, Tween, NP-40...). PEG contamination shows up in the mass spectrum as peaks separated by 44 Da. If detergents are present in your sample they will dramatically reduce the quality of your data.

NOTE: running the samples on a simple SDS PAGE gel will remove most PEG/PEG containing detergents from a sample.

2) Keratin is a common protein contaminant in a lot of proteomics samples. Keratin from your skin and hair is present in dust in the lab or can fall off you into your sample. It is important to try and minimise keratin contamination by working carefully and cleanly. Be aware that any surfaces, glassware or chemicals exposed to the lab atmosphere for more than a few minutes will be contaminated with keratin as dust from the atmosphere settles.

SOLUTION -

• Wash everything before use and then keep everything covered. Always close the caps on your microfuge tubes. Everything should be stored between uses in cupboards in sealed plastic bags or covered with aluminium foil.
• Always wear gloves when working with material for proteomics and work quickly. Working in a laminar flow hood will help to minimise dust and keratin but does not negate the need to be careful.
• When staining your gel do you leave the staining tray uncovered on the shaker? - cover the staining tray with aluminium foil at all times and wash the staining dish well before use.
• Common lab chemicals and materials, eg loading buffers, lab buffers, SDS, Gel tanks etc are common sources of keratin and dust. Use high quality and clean materials always. Use filtered deionized water always.
• DO NOT STORE ORGANIC SOLVENTS IN PLASTIC TUBES - contaminants leach out of the plastic with storage. Use new glass bottles or disposable glass scintillation vials

3) Polysiloxanes ([R2SiO]n) are a rare contaminant in proteomics samples and typically show up as peaks separated by 76 Da in the mass spectrum. Their source is typically from siliconized surfaces and plasticware where the coating is used to minimise adsorption of liquid to surfaces eg in high-recovery tips. Use high-quality, virgin plastic tips/tubes.

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These are expensive experiments. So please plan and execute experiments carefully.

a) Limit the number of samples to what is really necessary!!
b) Always use filtered deionized water!
c) Use nitrile (blue) gloves always!
d) Clean the whole gel running apparatus. Use fresh glove in all the steps and maintain clean surfaces throughout the process!!!!!
e) Use freshly prepared buffer and stains - do not re-use!
f) Always keep a printout of the gel picture with the marked gel slices!
g) Always use filtered deionized water!

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Do

• Fill out the request form in the portal and discuss your question before sending samples.
• Final sample submission is always by the PI.
• Provide the facility with as much information about your sample / protein as you can.
• Use clean containers to stain and store gels, and use them only for this purpose.
• Use a lid for the container mentioned above.
• Wear gloves at all times during sample handling.
• Cut gel bands on a clean bench and a clean glass plate using a clean (=new) scalpel knife.

DON'T

• DON'T incubate gels in the microwave to expedite coomassie staining
• DON'T use commercial kits for silver or coomassie staining without quality check
• DON'T use parafilm at all
• DON'T use overhead transparencies to scan gels. DON'T put the gel on scanner surface as well. Put the gel in clean transparent container and image it closed.
• DON'T use detergents for sample preparation in MS experiments of intact proteins
• NEVER use detergents to clean mass spec reagent bottles and other utensils.

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Protein identification by mass spectrometry is probabilistic, meaning that the best match is sought between an experimental spectrum and a theoretical spectrum for a peptide in the database. The score assigned to this match, and therefore the probability for the match to be right, depends on a number of parameters, such as spectral quality, mass accuracy, the size of the database, and the algorithm used for database searching. In addition, the more peptides are assigned to a given protein, the higher the protein score will be.

However, protein ID is based on peptides and there are many pitfalls that a result may be wrong, for example the same peptide is present in different proteins, or peptides from different proteins are taken together for a wrongly assigned protein.

We recommend that results have to be verified by independent experiments and the experiments have to be repeated.

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If you excised a single band from a gel, it is not unlikely that this contains several proteins of (almost) the same size, some of which may be below the detection level of the staining method used. Mass spectrometers by far exceed the sensitivity of the staining methods and it is normal to identify multiple proteins from single gel band.

For example, as much as 70 proteins from a single gel band can be expected when complex cell lysate is run on 12% minigel. It is better not to overload the gel as it affects the resolution and separation of proteins. Moreover, it can result in "carry over" of abundant peptides in LC resulting in identification of same protein repeatedly in different samples.

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There may be several reasons why this might occur.

• The instrument did not work properly. We take utmost care and run controls before running your samples. Still it may happen sometimes. We shall inform that to you immediately.
• Other reasons - the amount of starting material was simply not enough.
• Trypsin digestion did not work - always use high quality mass spectrometry grade trypsin.
• The protein under investigation does not contain (a sufficient number of) cleavage sites for trypsin. As a result, no peptides will be generated (and detected). This may be the case for some small proteins, but this may also occur to 'exotic' (e.g. highly acidic) proteins that contain fewer lysines and arginines than the average protein.
• The reverse might also be true: if a protein contains multiple cleavage sites, it will be digested into peptides that are too small to be detected, or to be sequenced with high confidence (e.g. histone tails). The alternative might be in the selection of another protease, which may be viable if you know the protein you are working with.
• Another reason for not finding a protein may be that it is not present in the database. This is not unusual for proteins originating from poorly characterized organisms.
• Finally, if you coomassie-stained your gel in a microwave or scanned on an overhead projector foil or prepared your samples in a dusty environment, these are the most likely reasons for not finding anything: microwaving bakes proteins in the gel, and there is no way to get them out, however intense the stain. Polymers in the overhead projector foils cause trypsin not to work. Dusty environment introduces contaminants which shadows the spectra of peptides from your sample leading to no identification.

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Yes, only if it is stained by mass spec compatible silver stained gel protocol. Many commercial vendors like Life technologies, Sigma, Pierce biotechnology (now Thermofisher) sell mass spec silver stain kits. However, before going for silver staining please consider the followings:

• Silver stain interferes with the MS analysis and requires more steps for sample preparation. In addition, the risk of contamination is increased during the long multi-step silver staining process.
• Silver staining also destroys proteins partially; the so called MS-compatible silver-stains are often only staining proteins on the surface of the gel, and they are not more sensitive than Colloidal Coomassie staining.

We always prefer Coomassie staining. If you are planning silver staining - inform the facility beforehand.

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For MALDI, we use the AB SCIEX Cal mix 5 for calibration. We also spot Beta-Galactosidase Digested from SCIEX and check its score and identification by protein Pilot soft ware.

The Orbitrao velos and Q-exactive is calibrated and tuned regularly. To assure the quality of the instruments, we run commercially available Standard Protein Digests before your samples.

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Yes, please ask for analysis help. We are planning to have training workshops at regular intervals. You are also welcome to ask for troubleshooting.

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We are planning to have training workshops at regular intervals. Please follow the portal for relevant notices.