Samples for Biotyping can be submitted Monday through Thursday by appointment only.
Protein samples are prone to degradation and must be analyzed as quickly as possible. Please email or call at least one day ahead of preparing the samples to confirm that the instrument and the operator will be available when your plate is ready.
The list of supplies and basic protocols for Biotyper can be found at the end of this post. In addition, the Biotyper manual in pdf format is available upon request. The Facility does not provide any of the supplies on this list and can only accept MS-ready target plates. For a demo on preparing the target plates, contact Tatiana.
To expedite your analysis, please fill out the Excel Sample ID Template with your sample names exactly as shown. All missing or incorrectly filled sample names will be replaced by the analysis date and your initials. You are responsible for keeping a record of your sample positions. The Excel file with the sample IDs must be emailed to Tatiana. Please attach any other information as separate files or documents.
Tips for successful microorganism identification
It is very important to exactly follow the procedure and use only the fresh, specified-grade reagents and solutions. MALDI is a competitive ionization process and is very sensitive to contaminants commonly present in histology-grade solvents and low-purity chemicals.
More is not better. A correct ratio of matrix to analyte is critical and achieved by following the procedure exactly.
Tube extraction (formic acid extraction) procedure requires extra few minutes of the prep time but yields much better results than the direct transfer procedure. Direct transfer procedure requires practice and is only applicable to non-spore-forming microorganisms. We recommend following the tube extraction procedure for all samples.
If you have any questions or suggestions or would like to discuss your project details, please email or call Tatiana.
Shipping the target plates
Plates can be shipped overnight on wet ice packs. To protect the sample spots, place the target plate facing down into its original box. You can tape it, so it does not get dislodged during shipping.
Electrospray ionization (ESI) and matrix-assisted laser desorption ionization (MALDI) are both competitive ionization processes in which a more abundant and/or more readily ionizable species will always win. Unless you are studying ionization of Tris, PEG, EDTA or CHAPS in the presence of small amounts of protein, the protein solution submitted for MS analysis should be essentially free of salts, detergents, chaotropic reagents, glycerol and other uninteresting interferences.
“So why not take a 10 mg/mL protein solution in a ‘mild buffer’ and dilute it to the required 0.1 mg/mL; the buffer concentration will be 100 times lower, and it is as good as desalted, right?” Wrong and here’s why:
Dilution is not a solution
Let’s say you have a 10 mg/mL solution of a 10 kDa protein in 10 mM Tris buffer, ‘mild’ enough. This means, you have a 10-fold molar excess of Tris compared to the protein. No matter how much you dilute this solution, the molar ratio of Tris to protein will remain 10:1 while it should be 1:50 or less.
For practical purposes, ‘essentially salt-free’ means that the total molar concentration of interfering species is much lower than that of the protein, e.g. at least 50-fold lower.
There are many ways to clean up a protein solution. In each case, the choice of a protocol will depend on the properties of interfering species and on the size and properties of the protein. For example, detergents can be removed using specialized detergent removal spin columns. PEGs and other charge-neutral contaminants can be washed away by loading the protein onto an ion exchange medium, eluting the protein with a high-salt buffer, and desalting the eluate. Buffer salts, urea, and other small molecules can be removed by either a desalting spin column packed with size-exclusion resin or a centrifugal filter with an appropriate molecular-weight-cut-off (MWCO) membrane. If you need an advice on your protein sample clean-up, I am always happy to help; you know where to find me!
Desalting using centrifugal filters
Centrifugal filters with different MWCO membranes are probably the easiest way to desalt a small volume of protein solution, although there are a few caveats. For example, the regenerated cellulose Ultracel membranes might retain cellulose-binding proteins; in this case, the low protein binding Omega membranes from PALL would be a better choice.
How many spin-dilute cycles are enough to ‘completely’ desalt a protein solution? This is one of those annoying situations where the old trusty triplicate doesn’t always work. Here’s my handy formula to help me decide when to stop desalting and start analyzing:
Let’s use our 10 mg/mL 10 kDa protein in 10 mM Tris buffer and a 3 kDa MWCO Amicon centrifugal filter as an example. The filter has a 0.5 mL capacity, and we can load 50 uL of the protein solution and dilute it with 450 uL of water. After a 30-min centrifugation, the volume is 50 uL, but the concentration of Tris is now 10% (or 0.1) of the initial while the protein concentration is essentially unchanged. So our D, dilution factor for Tris after each dilution+spin is 0.1; our goal, M final is 0.01 mM Tris (100:1 protein:Tris); and our M initial is 10 mM Tris. To find the number of dilute+spin cycles, dust off your scientific calculators:
Let’s throw in some urea, dilute the protein to 0.1 mM, and do our calculation again. How many steps (n = ?) do we need to get from 6 M to 1 uM urea (100:1 protein-to-urea ratio) using the same spin column as before?
You probably can get away with 6 dilute/spin repeats, but 7 would be better.
For those of you who hate equations, here’s a picture that illuminates the desalting process using the first example.
“What if my protein is unstable without a buffer?”
In that case, it may not be amenable to the direct ESI MS. However, before throwing up your hands, try to desalt your protein. It might be stable in water at micromolar concentration long enough to be analyzed – I am speaking from experience. You also have an option of using LC ESI MS or submitting your partially desalted protein for MALDI TOF MS.
No engineer and no news from the engineer. I have contacted Thermo.
SILAC: stable isotope labeling by amino acids in cell culture
SILAC incorporates stable-isotope labeled (or ‘heavy’) amino acids into cellular proteomes through normal metabolic processes. This is achieved by replacing natural (or ‘light’) amino acids in a growth medium with ‘heavy’ amino acids. The presence of heavy amino acids does not affect cell growth or morphology, but when ‘light’ and ‘heavy’ populations are mixed, the differences in protein abundances between the two populations can be determined by MS. SILAC affords the most accurate relative quantification compared with any chemical derivatization methods (e.g. iTRAQ or TMT) because the two (or more) samples are combined upstream of the protein extraction, at the intact cell level. Any cell culture system with defined amino acid sources is amenable to SILAC. Sufficient incorporation of ‘heavy’ amino acids usually requires several cell doublings; and the entire experiment can be completed within a few days.
Your first SILAC-based workflow should include a pilot experiment and a two-phase SILAC experiment. Once you are confident in your skills, your cells are adapted to the SILAC medium and have completely incorporated the ‘heavy’ amino acid(s), you can start with the part 2.2.
1. Pilot experiment (no labeling)
The purpose of a pilot experiment is to go through the entire procedure from growing cells to analyzing MS results, identify existing and potential problems, and find solutions prior to purchasing the isotopically-labeled amino acids. The pilot experiment is the only way to find out how many cells are needed to get good MS data, whether your enrichment/fractionation procedure is optimal, and whether the proteins of interest are well-represented.
The cost of MS analysis is low compared to the cost of isotopically labeled amino acids and the cost of other supplies and labor, so there is no advantage in shunning the ‘dry run’ experiment and moving on to the SILAC. The outcome of the pilot experiment is the optimized protocol.
2. SILAC experiment
2.1 Adaptation phase (untreated cells, heavy vs. light)
Adaptation phase experiment allows the cells to adapt to the SILAC medium and evaluates the degree of incorporation of the heavy amino acid over time. An aliquot of cells from only the heavy sample is analyzed. It is unnecessary to perform extensive fractionation of proteins before the analysis at this stage. A mixture of peptides from a gel band is analyzed by LC-MS, several peptides are selected, and the m/z region corresponding to the light form of each peptide is examined. In the final SILAC analysis, the proportion of proteins in the fully labeled ‘heavy’ state must be >98%, i.e. no detectable signal should be present in the m/z region of the ‘light’ form of a peptide.
2.2 Experiment phase (differential treatment, heavy vs. light)
Once the heavy amino acids are fully incorporated, two cell populations (from adaptation phase) can undergo a differential treatment, for example a drug is added to the ‘light’ population and the ‘heavy’ population is left untreated (control). The experiment follows the protocol that has been optimized during the pilot study. The outcome of the SILAC experiment is a record of quantitative and qualitative changes in the cell protein profile in response to the treatment.
A recent example of study that uses SILAC
‘Identifying novel targets of oncogenic EGF receptor signaling in lung cancer through global phosphoproteomics’, Zhang et al., Proteomics, 2015, 15, 340–355; doi: 10.1002/pmic.201400315
A generic SILAC protocol
‘A practical recipe for stable isotope labeling by amino acids in cell culture (SILAC)’, Ong and Mann, Nature Protocols, 2007, 1, 2650 – 2660; doi:10.1038/nprot.2006.427
SILAC methods, recipes, publications, links, etc.
The engineer called this afternoon; he is planning the service visit for Wednesday next week.
Orbi is down for an unknown time. I have placed a service call this afternoon and will keep you posted. The instrument went down in the middle of Joana’s 4th sample.
All submitted up to this point gel bands have been cleaned-up and are incubating with trypsin. I will lyophilize the peptides and store them in the freezer until the instrument is in operating condition.
Shhhh! Orbi is working… for now. Don’t spook it…
My standards ran 2 hrs ago. Gaozhong’s and Josh’s samples are on right now.
Orbi is still down, but I think I know why it shuts down. If I am unable to fix it today, I will place a service call. Katsu, Huihun, and Nick – your samples ran last night, and I will send you the data tomorrow. Joana, only first set of your samples is done; second set is ready to go as soon as Orbi is working again. Everyone else – please be patient!
Orbi is not ready, it vented for an unknown reason some time last night. I turned it on again this morning, and now we wait and hope that the ghost in the machine is in a better mood today.