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When you place an order, a link with shipping instructions, and shipping labels, if requested, will appear on the order page. Please read the shipping instructions carefully for information on how to label your samples and how to pack and ship your samples. The instructions contain a packing list of two or more pages which must be included in the shipment.
A brief overview of some sample-specific requirements are listed here. Sample preparation is included in the price of our assays. More generally, sample preparation should be minimal. We have the equipment and the expertise to process all common sample matrices, including affinity purifications, biofluids, adherent and suspension cell cultures, conditioned media, host-cell protein isolates, animal tissues, plant tissues, as well as food and beverage products, supplements and cosmetics. We can pulverize liquid nitrogen-frozen samples, bead beat yeast, fungi or bacteria, and probe sonicate. From there we can extract and isolate protein away from particulates and cellular components such as lipids and nucleic acids.
We can clean up samples containing almost all commonly used buffers and detergents, including SDS, Triton, Tween and many others. When placing an order simply indicate the known buffer components in the sample description, or if unknown, the catalog numbers of the kits used to prepare the samples. When we review your order we'll let you know if there are any issues.
Sample quantity recommendations for common sample matrices are provided here. In general, for profiling assays we are looking for 1-10s of micrograms of total protein. For phosphorylation assays, 100s-1000s of micrograms of total proteins.
Modern mass spectrometers are very sensitive, with routine limits of detection at the mid- to low-attomole level of protein digest loaded "on-column." Bear in mind that the sample we receive is not loaded "on-column" in its entirety. Depending on the sample amount and matrix we may extract protein from all or a portion of the sample, digest all or a portion of the extracted protein and load all or a portion of the digest. There is some sample loss during sample preparation. We generally try to keep some sample in reserve to allow multiple analyses if necessary, but that is not always possible.
In general, the best way to evaluate a sample is to run it on SDS-PAGE. Extract protein by boiling the sample in 10% SDS. Depending on the sample matrix you may need to mechanically homogenize the sample first. Spin out any debris and follow the instructions in your SDS-PAGE kit for sample preparation and loading to the gel. Run the gel, then stain with CBB or silver. If you can see bands there is enough protein in the sample for us to work with.
Visualization by Western blot is not a good indicator of protein quantity and cannot be reliably used to predict detection by LCMS. Make sure you can see the band of interest by CBB or silver stain.
Another consideration is that only a fixed amount of sample can be loaded "on-column." For nano-scale LC this is typically on the order of about one microgram of protein digest. If your protein is present below the limit of detection based on that load amount, your protein will not be detectable by the instrument. This is perhaps best exemplified by plasma and serum, which are one of the most complex sample types, and also have the highest dynamic range of protein abundance. Albumin and IgGs alone constitute about 2/3 of the total protein mass, while many biomarkers are present at low pg/mL levels. The 10 most abundant proteins constitute over 90% of the protein mass. This effectively "limits" the amount of other protein which is loaded "on-column."
There are a couple of commonly utilized workarounds for this limitation. Immunoaffinity columns which target albumin, IgG and other abundant proteins are often used for plasma and serum samples to deplete those abundant proteins, effectively enriching the lower abundance proteins in the sample. Fractionation is commonly used for other sample types to split the protein digest into 2 or more fractions which are then analyzed separately. because one microgram of fractionated sample can be loaded on-column, the effective protein load is equal to one microgram multiplied by the number of fractions. The drawback to this method is that it increases analysis time linearly by the number of fractions analyzed.
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Yes, Plus users are able to upload custom protein sequence files in FASTA format. The FASTA files are automatically cleaned, validated and made otherwise suitable for use in our data processing pipelines. Each custom FASTA file can be up to 1.5 GB. FASTAs are securely stored and permanently associated with the project. Any of your custom FASTAs can be used in other projects.
Why are custom FASTAs only available to Plus and Pro customers?
These plans provide extended support services, as well as long-term data access and storage. Custom FASTAs often require additional support and long-term access.
A few micrograms of peptide digest is typically loaded for an assay. To account for sample losses and to permit re-analysis if necessary, we request about 50 micrograms of total protein per assay. Good results can be obtained from trace sample amounts - we can identify over 1,000 proteins from less than 1,000 cells procured via laser dissection.
Please send in the smallest volume possible. We will acetone precipitate protein in sample volumes greater than 2 mL.
Any species with an available sequence library (FASTA file) can be assayed. If a sequence library is not available for a species or if the library is limited in scope, we can attempt to search a library of related species to search for homologous proteins. We support NCBI and UniProt libraries by default. Search UniProt taxonomies to see if your organism is present. Subscribers to our "Plus" support plan can upload custom sequence libraries as FASTA files.
In a typical complex sample such as a cell line we identify, on average, 5 different peptide sequences mapping to each protein and an average of 2 peptide spectral matches (identification events) per peptide.
The Q-Exactive has a sub-femtomole limit of detection (LOD). The LOD, limit of identification and limit of quantification for each peptide will vary based on several factors such as ionization efficiency and fragmentation characteristics.
We prefer SILAC, but it is limited to cell culture experiments. SILAC controls for variability throughout the sample preparation and UPLC-MS/MS assay. In contrast, stable isotope dimethyl labeling can be applied to any sample type post-digestion. Because of this, it only controls for variability post-derivitization - primarily for UPLC-MS/MS.
Another option is to use a super-SILAC spike-in, in which a set of SILAC cultures are mixed to provide an adequate representation of the proteome. This super-SILAC mix is then mixed with the sample. This also controls for variability throughout sample preparation and UPLC-MS/MS.
We do not support isobaric tagging experiments (iTRAQ or TMT). We feel the instrumentation is not yet advanced enough to perform these assays without serious compromises which render this type of experiment less effective than label-free, SILAC or stable isotope dimethyl labeling experiments.
You will receive a free account on Proteome Cluster for viewing all details of the assays, including protein and peptide assignments, tandem mass spectra, run-level metrics, search parameters, etc. All data is available to download or share with your colleagues and is stored indefinitely.
Yes, we offer both iron- and titanium oxide-based phoshpopeptide enrichments. Due to the very low amount of phosphopeptides present in a sample, we recommend providing at least 1-10 mg of total protein. This will permit the identification of thousands of discrete phosphopeptides.
We also offer lectin affinity-based glycopeptide or glycoprotein enrichments.
Yes, we offer albumin, immunoglobulin and multi-protein depletions from fluids such as plasma, urine and saliva.
In a nutshell, proteins are digested to peptides which are separated by liquid chromatography and ionized for introduction into a mass spectrometer where they are fragmented in a process call tandem mass spectrometry. The resulting fragmentation pattern is compared to theoretical fragmentation patterns of peptides of similar mass derived from a protein sequence library and scored based on the quality of the match. A single match can be sufficient to confidently identify a protein and categorize it as present in the sample. Multiple matches to multiple peptide sequences from the same protein sequence can, in aggregate, be used to validate that a protein with the predicted protein sequence is present in the sample.
High-performance liquid chromatography (HPLC) is a method used to separate analytes according to one or more properties of the analytes. In the case of shotgun proteomics, tryptic peptides are most often separated based on their hydrophobicity using a reversed phase column. This is done to both slow the entry of peptides in the mass spectrometer and to maximize the concentration of each peptide as it enters the mass spectrometer by separating them from each other. A mass spectrometer can only sample a finite number of peptides in a unit time (about 10 per second on a Thermo Q-Exactive Orbitrap). Separation of the peptides is accomplished by running an increasing concentration of acetonitrile through the column. As the concentration of acetonitrile increases, increasingly hydrophobic peptides will elute from the column for introduction into the mass spectrometer.
Nanoflow liquid chromatography (nLC) can improve overall sensitivity by decreasing the volume in which a peptide elutes, thereby increasing the concentration of the peptide per unit time as it enters the mass spectrometer. Running nLC involves using small diameter columns (typically 50 or 75 micron inner diameter) and nanoliter per minute flow rates (typically 100 - 300 nanoliters per minute) and requires the use of an HPLC capable of pumping a gradient at nanoliter per minute flow rates Thermo Easy-nLC II or 1000).
Longer chromatographic columns and smaller diameter chromatographic media coupled with nLC can further improve chromatographic resolution, which increases peptide concentration per unit time, resulting in increased sensitivity. However, longer columns and smaller diameter media require increased pressure to pump the separation gradient through the column. Ultra-high pressure chromatography (UPLC) (Thermo Easy-nLC 1000) can deliver the pressure necessary to perform separations on 50 cm columns with 2-micron media. This enables highly resolving separations of 4 hours or more, negating, for many applications, the need for pre-fractionation of the peptides prior to UPLC.
Peptides must be transferred from liquid- to gas-phase for introduction to a mass spectrometer. There are numerous ionization methods, but the two most commonly used utilize either a laser or an electric current for the transfer. These methods shared the 2002 award for the Nobel Prize in Chemistry.
Tandem mass spectrometry is the process by which a peptide is isolated, fragmented and detected inside a mass spectrometer. Once a mass spectrometer detects the presence of a peptide (typically as a high intensity, multiply-charged ion) a decision is made to isolate the ion within a defined mass-to-charge (m/z) window. Isolation can be done “in time” (on time-of-flight devices) or “in space” (on trap devices). Once isolated, the peptide is fragmented. There are numerous fragmentation methods. Two of the more common methods are termed collision induced dissociation (CID, alternately collision activated dissociation or CAD) and higher energy collisional dissociation (HCD). Both methods fragment a peptide primarily along the peptide backbone, generating a population of peptide ions with one or more amino acids cleaved from either end. Peptide ions with amino acids removed from the C-terminus and N-terminus are denoted b-ions and y-ions, respectively. CID typically yields both b- and y-ions, potentially allowing for the peptide to be sequenced from both ends. HCD typically yields primarily y-ions as well as immonium ions in the low mass region.
Shotgun proteomic search algorithms take as input a peak list file containing data from a single tandem mass spectrum including: the precursor (intact) mass of the peptide isolated for fragmentation, the charge state of the peptide and the masses and intensities of each of the fragment ions. The algorithm takes the protein sequence library defined in the search (typically the entire sequence library for a species such as Homo sapiens) and performs an "*in silico*" digestion to generate all possible peptides with tryptic ends. From this set of peptides a subset is selected which have a precursor mass within the mass tolerance range defined in the search algorithm parameters (typically +/- 1.5 Da on on ion trap and +/- 20 ppm on an Orbitrap). The algorithm then generates theoretical fragmentation patterns for each peptide in this subset, compares the experimentally-obtained fragmentation spectrum to each of these and assigns a score to each. Various methods are used to estimate the likelihood that the highest scoring match is a random event. Some algorithms, such as OMSSA and X!Tandem, calculate expectation values. For these, an expectation value score of 0.01 indicates that there is a 1 in 100 chance that the match is a random event. In other words, in a population of 100 matches with expectation value scores of 0.01 we would expect that one of the matches would be a random event, that is, a false positive identification, but we would not know which one.
A shotgun proteomics search algorithm uses two mass tolerance filters. One for the precursor ion mass measurement (intact peptide mass) and another for the fragment ion mass measurements. For measurements with higher mass accuracy these tolerances may be tightened, such that there are fewer candidates for a match. This results in a higher score for a given tandem mass spectrum, decreasing the likelihood of a false positive assignment. As mass accuracy increases and tighter mass tolerances are used, fewer theoretical mass spectra need to be evaluated by the algorithm which decreases the amount of time necessary for the algorithm to complete. An ion trap mass spectrometer typically has nominal mass accuracy. That is, a mass assignment is accurate to within about +/- 1 Da. An Orbitrap mass spectrometer has a mass accuracy of better than +/- 10 ppm. The difference between the two is the capability of being able or not being able to distinguish glutamine from lysine, a peptide from its deamidated counterpart, and a tri-methylation modification from an acetylation modification.
Increasing mass resolution enables increasing capability to determine the charge state of a peptide in shotgun proteomics experiments by resolving the isotopic envelope. This allows the set of theoretical spectrum candidate matches to be narrowed, increasing the resulting score and decreasing the likelihood of a false positive match, similar to better mass accuracy. Increasing mass resolution also allows for improved calculation of precursor peak areas by resolving the peaks of peptides with similar masses.
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