In the SMOAC protocol (https://www.thermofisher.com/blog/learning-at-the-bench/wp-content/uploads/sites/13/2024/10/High-SelectTM-SMOAC-Protocol.pdf), can I enrich with High-Select Fe-NTA kit first?
No. It is important to enrich with the TiO2 kit (Cat. No. A32993) first. Afterwards, the flow-through and wash fractions from this enrichment can be processed with the Fe-NTA kit (Cat. No. A32992). If this order is reversed (that is, Fe-NTA before TiO2), there will be 2 consequences as follows:
1. There will not be any significant additional recovery of peptides (maybe just a few more peptides).
2. There will be no enrichment for the multiple phosphorylated peptides, so those would be lost.
Why do you offer two phosphopeptide enrichment kits: the High-Select TiO2 kit (Cat. No. A32993) and the High-Select Fe-NTA kit (Cat. No. A32993)?
The two phosphopeptides enrichment kits, Fe-NTA and TiO2, enrich a complementary set of phosphopeptides.
Our R&D has developed a Sequential enrichment of Metal Oxide Affinity Chromatography (see https://assets.thermofisher.com/TFS-Assets/CMD/posters/PO-65032-SMOAC-Phosphoproteomics-Peptides-ASMS2017-PO65032-EN.pdf and https://www.thermofisher.com/blog/learning-at-the-bench/wp-content/uploads/sites/13/2024/10/High-SelectTM-SMOAC-Protocol.pdf?CID=bid_mic_r04_jp_cp0000_pjt0000_bid00000_0so_blg_protein_analysis_mass_spectrometry_bid_ts_mbr_24065_Social_LAB) where flow-through and wash fractions from TiO2 enrichment were combined and subjected to Fe-NTA enrichment. This sequential enrichment provides impressive coverage of phosphoproteomes.
What are the differences between the old Fe-NTA kit (Cat. No. 88300) and the new High-Select Fe-NTA kit (Cat. No. A32992)?
There are four differences between the Fe-NTA kit (Cat. No. 88300) and the new High-Select Fe-NTA kit (Cat. No. A32992) kit as follows:
1. The selectivity - the ratio of number of phosphopeptides over total peptides - was significantly improved to 99% with Cat. No. A32992, because the reagents were extensively optimized for the phosphopeptide selection.
2. The phosphopeptide yield was also increased to 33 µg based on quantitative colorimetric peptide assay (Cat. No. 23275).
3. The reagent is a pre-formulated format, so mixing reagent to prepare the working solution from stock solutions provided in the old kit (Cat. No. 88300) is not necessary, so it is easier to handle.
4. The enrichment protocol is optimized and streamlined, which means there are many fewer steps than with Cat. No. 88300. Thus, it takes <45 min to finish the entire protocol compared to 2 hours with the old kit (Cat. No. 88300).
What is the standard solvent for the Pierce Quantitative Peptide Assays & Standards (Cat. No. 23275)?
The solvent for the Pierce Quantitative Peptide Assays & Standards (Cat. No. 23275) is 50mM ammonium bicarbonate.
When I quantitate my mass spec peptide sample with the Pierce Quantitative Colorimetric Peptide Assay, I get different results than when I use the Pierce Quantitative Fluorometric Peptide Assay. Which is best to use for the most accurate quantitation?
Since the different peptide assays use different chemistries to measure peptides, they may result in different results. Interfering compounds are the most common source of background and inaccurate measurements. Please note that the fluorometric peptide assay is not recommended for peptides which have been modified using TMT reagents.
How can I determine peptide yield in my mass spectrometry samples after sample clean-up?
Peptide yield can be measured using the Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275) or the Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290). The choice of peptide assay depends on the sample type and composition of the sample buffer. The fluorometric peptide assay cannot be used to measure peptides with chemically modified amines such as acetylated peptides or TMT-labeled protein digests. The colorimetric assay can measure a wider range of samples but is not as sensitive as the fluorometric assay, requiring more sample for accurate detection. Finally, both assays are susceptible to interfering compounds in the sample or buffer which should be avoided or removed for best results.
I have good peptide identification numbers but the variation between sample replicates is high. What do you recommend to improve sample reproducibility?
We recommend reviewing your sample-prep workflow to ensure consistent protein extraction, reduction/alkylation, digestion, and clean-up. We recommend using EasyPep products for high quality, reproducible sample preparation (EasyPep Maxi Sample Prep Kit, EasyPep Mini MS Sample Prep Kit, EasyPep 96 MS Sample Prep Kit). We also recommend quantifying peptides using the Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290) or Pierce Quantitative Coloimetric Peptide Assay (Cat. No. 23275) to ensure that the same amount of peptides are being used for each LC-MS analysis. Poor reproducibility could also be related to the LC-MS system performance which may require recalibration using Pierce Calibration Solutions. System performance can be assessed using protein digest standards such as Pierce HeLa Protein Digest Standard or Pierce TMT11plex Yeast Digest Standard and peptide standards such as Pierce Peptide Retention Time Calibration Mixture or Pierce LC-MS/MS System Suitability Standard (7 x 5 Mix).
I have varying amounts of peptide in my samples. Is there a way to determine peptide yield after sample clean-up?
Peptide concentration can be measured using our Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290) or Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275).
I have varying amounts of peptide in my mass spectrometry analysis samples. Is there a way I can determine peptide yield after sample clean-up?
You can measure peptide concentration using our Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275) or Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290).
My buffer or components of my buffer are not listed in the compatibility table for my protein assay. What should I do?
You can test the tolerance of the assay for your specific buffer formulation. For in-house generated compatibility information, substances were considered compatible at the indicated concentration in the Standard Test Tube Protocol (found in the manual for each protein assay) if the error in protein concentration estimation caused by the presence of the substance was less than or equal to 10%. The substances were tested using WR prepared immediately before each experiment. Blank-corrected 562nm absorbance measurements (for a 1000µg/mL BSA standard + substance) were compared to the net 562nm measurements of the same standard prepared in 0.9% saline.
All the components of my sample buffer are at or below the indicated compatible concentration for my protein assay, but I am still seeing too much/too little color development. What could be the problem?
It is possible to have a substance additive affect such that even though a single component is present at a concentration below its listed compatibility, a sample buffer containing a combination of substances could interfere with the assay. You should take steps to eliminate or minimize the effects of the interfering substance(s) by diluting or removing the substance.
My protein assay is not developing color or is developing too much color. What can I do?
Refer to the information in the product-specific instruction booklet or our Tech Tip: Protein Quantitation Assay Compatibility Table (https://assets.thermofisher.com/TFS-Assets/LSG/Application-Notes/TR0068-Protein-assay-compatibility.pdf).
My spectrophotometer doesn’t have a filter set for the absorbance maximum. Can I use an alternate wavelength to read the protein assay?
Often, an alternative wavelength can be used, although the slope of the standard curve and the overall assay sensitivity will most likely be reduced. Our Tech Tip (https://tools.thermofisher.com/content/sfs/brochures/TR0025-Protein-assay-spectra.pdf) offers additional information on determining acceptable wavelengths for measuring protein assays.
What other factors affect the protein assay accuracy and precision?
Several factors affect protein assay accuracy and precision:
Replicates: The only way to evaluate the extent of random error is to include replicates of each standard and test sample. Because all test samples are evaluated by comparison to the standard curve, it is especially important to run the standards in at least triplicate. The standard deviation (SD) and coefficient of variation (CV) can then be calculated, providing a degree of confidence in your pipetting precision. If replicates are used, curve-fitting is done with the average values (minus obvious outliers).
Blank correction: It is common practice to subtract the absorbance of the zero assay standard(s) from the all other sample absorbance values. However, if replicate zero-assay standards will be used to calculate error statistics, then another independent value may be required for blank-correction. If the standards were prepared in a buffer to match that of the test samples, and this buffer contains components that may interfere with the assay chemistry, it is informative to blank the absorbances with a "water reference" (i.e., a zero-protein, water sample). Differences between the water reference and zero standard sample are then indicative of buffer effects.
Standard curve slope: The standard curve slope is directly related to assay accuracy and sensitivity. All else being equal, the steepest part of the curve is the most reliable. For most protein assays, the standard curve is steepest (i.e., has the greatest positive slope) in the bottom half of the assay range. In fact, the upper limit of an assay range is determined by the point at which the slope approaches zero; the line there is so flat that even a tiny difference in measured absorbance translates to a large difference in calculated concentration.
Measurement wavelength: The measurement wavelengths that are recommended for each protein assay method are optimal because they yield standard curves with maximal slope. This usually, but not always, corresponds to the absorbance maximum. (In certain circumstances, other considerations are also important in choosing the best possible measurement wavelength, such as avoiding interference from sample components that absorb at similar wavelengths). In fact, for most protein assays, depending on the precision required, acceptable results can be obtained using any measurement wavelengths within a certain range.
When does a dilution factor need to be applied in a protein assay?
One situation in which the dilution factor is important to consider is when the original sample has been pre-diluted relative to the standard sample. Suppose the original protein sample is actually known to be approximately 5 mg/mL. This is too concentrated to be assayed by the Pierce Bradford Plus Protein Assay Kit, for example, whose assay range in the standard microplate protocol is 100-1500 µg/mL. However, you could dilute it 5-fold in buffer (i.e., 1 part sample plus 4 parts buffer) and then use that diluted sample as the test sample in the protein assay. If the test sample produces the same absorbance as the 1000 µg/mL standard sample, then you can conclude that the test (5-fold diluted) sample is 1000 µg/mL, and therefore the original (undiluted) sample is 5 x 1000 µg/mL = 5000 µg/mL = 5 mg/mL.
Do I need to know the protein concentration in the assay reagent for my protein assay?
No. It is neither necessary nor helpful to know the protein concentration as it exists when the samples are diluted in assay reagent. The protein concentration when diluted by assay reagent is almost certainly not the value of interest; instead, one wants to know the protein concentration of the original test sample.
Do I need to know the amount of protein per well for my protein assay?
No. Contrary to what many people assume, it is neither necessary nor even helpful to know the actual amount (e.g., micrograms) of protein applied to each well or cuvette of the assay. The amount of protein per well is almost certainly not the value of interest; instead, one usually wants to know the protein concentration of the original test sample.
How can I utilize Excel software to plot and apply the standard curve for my protein assay?
Enter the concentration values for the standards in Column A and their corresponding absorbance data in Column B. Highlight both columns and from the Insert menu select Chart and XY (Scatter). Click on the resulting graph and select Add Trendline from the Chart menu. While viewing the graph next to the open Format Trendline window, choose Polynomial and set the Order to 2, 3 or 4 until the best-fit appears. Check the box near the bottom called Display Equation on Chart; then close the Format Trendline window. Use the resulting equation to determine protein concentration (y) of an unknown sample by inserting the sample’s absorbance value (x).
How can I interpolate my protein assay data?
Most modern plate readers and spectrophotometers have associated software that automatically plots a linear or curvilinear regression line through the standard points, interpolates the test samples on that regression line, and reports the calculated value. However, there are different methods for making the calculations “by hand”. You can find a detailed explanation and example in our Tech Tip.
How can I accurately analyze my protein assay data?
With most protein assays, sample protein concentrations are determined by comparing their assay responses to that of a dilution-series of standards whose concentrations are known. The responses of the standards are used to plot or calculate a standard curve. Absorbance values of unknown samples are then interpolated onto the plot or formula for the standard curve to determine their concentrations. The most accurate results are possible only when unknown and standard samples are treated identically. This includes assaying them at the same time and in the same buffer conditions, if possible. Because different pipetting steps are involved, replicates are necessary if you wish to calculate statistics (e.g., standard deviation, coefficient of variation) to account for random error. It is imperative to run a new standard curve for each set of samples to be tested
It was necessary to dilute my sample in order to run the protein assay (i.e,. due to an incompatible substance). How do I account for this when determining the concentration?
Simply multiply the calculated concentration of the diluted sample by the dilution factor. For example: A protein sample is known to be approximately 5 mg/mL. This is too concentrated to be assayed by the Pierce Bradford Plus Protein Assay Kit, whose assay range in the standard microplate protocol is 100-1500 µg/mL. However, you could dilute it 5-fold in buffer (i.e., 1 part sample plus 4 parts buffer) and then use that diluted sample as the test sample in the protein assay. If the test sample produces the same absorbance as the 1000 µg/mL standard sample, then you can conclude that the test (5-fold diluted) sample is 1000 µg/mL, and therefore the original (undiluted) sample is 5 × 1000 µg/mL = 5000 µg/mL = 5 mg/mL.
In my protein assay, what unit of measurement will my sample concentration be in after calculating the concentration?
The unit of measure used to express the standards is by definition the same unit of measure associated with the calculated value for the unknown sample (i.e., final results for unknown samples will be expressed in the same unit of measure as was used for the standards). For example, if the standard concentrations are expressed as micrograms per milliliter, then the concentrations for the unknown samples, which are determined by comparison to the standard curve, are also expressed as micrograms per milliliter.
What should I dilute my protein standard in for my protein assay?
Protein standards should preferably be diluted using the same diluent as the sample(s). Sample assay responses are directly comparable to each other if they are processed in exactly the same manner. Variance in protein quantity is the only possible cause for differences in final absorbance (color intensity) if samples are dissolved in the same buffer and the same stock solution of assay reagent is used for all samples.
However, if only a “rough” estimate of protein concentration is needed, a blank-only correction can be used. In this case, a blank is prepared in the diluent of the sample to correct for its raw absorbance. The concentration of the sample is then determined from a standard curve obtained from a series of dilutions of the protein of known concentration prepared in water or saline solution.
What protein should I use to generate my standard curve?
Protein concentrations are generally determined and reported with reference to standards of a common protein, such as bovine serum albumin (BSA). If precise quantitation of an unknown protein is required, it is advisable to select a protein standard that is similar in quality to the unknown; for example, a bovine gamma globulin (BGG) standard may be used when assaying immunoglobulin samples.
Why is the choice of protein standard important in a protein assay?
Because proteins differ in their amino acid compositions, each one responds somewhat differently in each type of protein assay. Therefore, the best choice for a reference standard is a purified, known concentration of the most abundant protein in the samples. This is usually not possible to achieve, and it is seldom convenient or necessary. If a highly purified version of the protein of interest is not available or it is too expensive to use as the standard, the alternative is to choose a protein that will produce a very similar color response curve in the selected protein assay method and is readily available to any laboratory at any time. Generally, bovine serum albumin (BSA) works well as a protein standard because it is widely available in high purity and relatively inexpensive. Alternatively, bovine gamma globulin (BGG) is a good standard when determining the concentration of antibodies because BGG produces a color response curve that is very similar to that of immunoglobulin G (IgG).
What are the basic principles of standard curve assays?
- Identically assayed samples are directly comparable: Sample assay responses are directly comparable to each other if they are processed in exactly the same manner. Variation in amount of protein is the only possible cause for differences in final absorbance (color intensity) if the samples are dissolved in the same buffer, the same lot and stock solution of assay reagent is used, all samples are mixed and incubated at the same time and temperature, and no pipetting errors were introduced.
- Units in equals the units out: The unit of measure used to express the standards is by definition the same unit of measure associated with the calculated value for the unknown sample (i.e., final results for unknown samples will be expressed in the same unit of measure as was used for the standards).
What is protein-to-protein variation?
Each protein in a sample responds uniquely in a given protein assay, and this protein-to-protein variation is observed as differences in the amount of color (absorbance) obtained when the same mass of various proteins is assayed concurrently by the same method. These differences in color response relate to differences in amino acid sequence, isoelectric point (pI), secondary structure, and the presence of certain side chains or prosthetic groups.
Depending on the sample type and purpose for performing an assay, protein-to-protein variation is an important consideration in selecting a protein assay method and in selecting an appropriate assay standard (e.g., BSA vs. BGG). Protein assay methods based on similar chemistry have similar protein-to-protein variation.
How should a sample be prepared before a protein assay?
Before the sample is analyzed, it must be solubilized in a buffered aqueous solution. Depending on the source material and the procedures involved before performing the protein assay, the sample will likely contain a variety of non-protein components. Awareness of these components is critical for choosing an appropriate assay method and evaluating the cause of anomalous results. Every type of protein assay is adversely affected by substances of one sort or another. Components of a protein solution are considered interfering substances in a protein assay if they artificially suppress the response, enhance the response, or cause elevated background by an arbitrarily chosen degree (e.g., 10% compared to control). Additional components can include reducing agents, chelators, crowding agents, and protease inhibitors.
What should I consider when choosing a protein assay?
There are several criteria that should be considered, including compatibility with the sample type and components, assay range and required sample volume, protein-to-protein uniformity, speed and convenience for the number of samples to be tested, and the availability of spectrophotometer or plate reader necessary to measure the color produced (absorbance) by the assay.
Why does the protein assay method matter?
Unfortunately, no protein assay method exists that is either perfectly specific to proteins (i.e., not affected by any nonprotein components) or uniformly sensitive to all protein types (i.e., not affected by differences in protein composition). Therefore, successful use of protein assays involves selecting the method that is most compatible with the samples to be analyzed, choosing an appropriate assay standard, and understanding and controlling the particular assumptions and limitations that remain. The objective is to select a method that requires the least manipulation or pre-treatment of the samples to accommodate substances that interfere with the assay. Each method has its particular advantages and disadvantages. Because no one reagent can be considered the ideal or best protein assay method for all circumstances, most researchers have more than one type of protein assay available in their laboratories.
What protein assay is best?
Unfortunately, no protein assay method exists that isn’t affected by any non-protein component or uniformly sensitive to all protein types. One must select an appropriate assay method based on compatibility with the sample type or one that requires the least manipulation of the sample to accommodate the assay. Most researchers will have more than one type of assay available in their laboratories.
I need to measure the concentration of a peptide, which protein assay can I use?
Typically, peptides need to be around 2 or 3 kDa (depending on the protein assay and the exact peptide composition) to be measured using a protein assay. For peptides, we offer two quantitative assays, the Quantitative Fluorometric Peptide Assay (Cat. No. 23290) and the Quantitative Colorimetric Peptide Assay (Cat. No. 23275).
What protein assays do you offer for total protein quantitation?
We offer several types of protein assays including the: BCA Assay, BCA-RAC (Reducing Agent Compatible) Assay, Micro BCA Assay, 660 nm Protein Assay, Pierce Bradford Plus Protein Assay Kit, Pierce Bradford Protein Assay Kit, Modified Lowry Assay, colorimetric and fluorometric Peptide Assays, CBQCA kit, EZQ kit, Quant-iT kits, NanoOrange, and the Qubit kits.
