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View additional product information for Qubit™ Protein and Protein Broad Range (BR) Assay Kits - FAQs (A50668, A50669, Q33212, Q33211)
39 product FAQs found
The accuracy and sensitivity of the Qubit quantitation assays are the same as that of a microplate reader. This was a requirement during product development. The detection limits for each Qubit kit can be found on the corresponding product manual, which can be found by searching our website by keyword or catalog number.
Find additional tips, troubleshooting help, and resources within our Nucleic Acid Quantification Support Center.
No. The Qubit DNA and RNA kits only quantify the amount of either DNA or RNA in the sample. The Qubit fluorometer cannot take absorbance readings to provide a A260/A280 ratio or detect protein in nucleic acid samples. This can be done with the NanoDrop instrument. If your sample contains protein or other contaminants that can affect the assay, it should be further purified.
If your sample may contain both DNA and RNA, one may use either (or both) the DNA and RNA Qubit kits and compare with samples treated with either RNase or DNase to get an accurate determination of DNA or RNA, respectively.
All Quant-iT and Qubit kits are compatible with all fluorometers and microplate readers that have the appropriate light sources and filters. You won't have access to the algorithm in the Qubit fluorometer for generating the standard curve provided by the instrument, instead, you must make a few dilutions of the highest standard DNA or RNA (Standard #2) in the Qubit kits to generate a standard curve with multiple data points.
No, we do not recommend this. Some of the dyes in the original Quant-iT kits (those NOT listed as “for use with the Qubit fluorometer”) are not compatible with the Qubit Fluorometer. In addition, the new Quant-iT kits (labeled as “for use with the Qubit Fluorometer”) have standards formulated to be compatible with the Qubit fluorometer internal algorithms for the respective assays. The Qubit Fluorometer-compatible kits are also less expensive per assay if you are processing fewer than 20 samples at a time.
The concentration of detergents in 1x RIPA buffer is quite high, therefore, we do not recommend that you use this kit with samples in RIPA. The Qubit Protein Assay has a tolerance for detergents present only in very low amounts. Please see the user guide for the tolerance of this assay for detergents here.
The Qubit Protein Broad Range (BR) Assay Kit (A50668) has a much higher tolerance for detergents, e.g. RIPA buffer can be used undiluted. Please see this link.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Quantitation Support Center.
To be able to use the Qubit Protein Broad Range Assay (Cat. No. A50668) on the Qubit 4 Fluorometer (Cat. No Q33238) you would need to install the latest firmware on the fluorometer. Please find the latest Qubit 4 firmware (v2.19) on the following webpage: https://downloads.thermofisher.com/Qubit4/v2.19/Qubit4_update_v2.19.pak
Save the firmware update on a USB stick and upload it to the Qubit 4 Fluorometer.
Once the firmware update has been installed, you will be able to see the Protein BR assay on the Qubit 4 Fluorometer.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Quantitation Support Center.
Though the Quant-iT Protein Assay Kit is indicated for high-throughput application, it is completely separate from the Qubit Protein and Protein Broad Range Assay Kits, not another version of them.
No, the Quant-it and Qubit reagents use different dyes and therefore are not interchangeable.
To access “Protein Broad Range” in the “Protein” menu of your Qubit 4 instrument, as instructed in the Qubit Protein BR Assay Kit (Cat. Nos. A50668, Q33211, Q33212, A50669) instructions, the instrument will require the latest firmware update (v2.19 or v1.8.0 for the Qubit Flex). You can check which version you currently have on your device by clicking on "Settings" from the Home screen, and then on "About Instrument".
To update the software:
Download the latest firmware update from the Qubit Fluorometers Technical Resources page.
Follow the instructions for updating your Qubit instrument in the firmware Release notes or in the Qubit 4 Fluorometer User Guide (page 53).
After the update has been completed, the option "Protein Broad Range" should become available.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Quantitation Support Center.
The Qubit Protein Assay is compatible with very small amounts of detergent. See “Contaminants Tolerated by the Qubit Protein Assay” Table 2 on page 6 of the Qubit Protein Assay Kit product manual (https://tools.thermofisher.com/content/sfs/manuals/Qubit_Protein_Assay_UG.pdf) for specific amounts.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
Here are possible causes and solutions:
- The kit has expired or has been stored incorrectly: When properly stored, the components of the Qubit Protein Assay Kit should be stable for at least 6 months. Upon receipt, the kit can be stored at 4 degrees C. Components A and B can be stored at ambient temperature and Components C-E can be stored at ≤4 degrees C. Protect Component A from light. High degradation of the BSA standard 2 and 3 will result in a decrease in signal and a “Standards Incorrect” error warning upon calibration. Replace the kit.
- Old calibration data was used: Best practice is to prepare fresh calibration standards at the same time as the samples to take into account any changes in assay conditions.
- Incorrect tubes were used: Use the recommended Qubit Assay Tubes (Cat. No. Q32856). Other 0.5 mL thin-walled PCR tubes may work as well, but performance is not guaranteed. Avoid opaque tubes, as these will block the light path.
- Tubes contain bubbles or particulates that are scattering the light: Pipette gently to avoid the introduction of bubbles. Spin down tubes before measuring to remove bubbles or particulates. Spin down samples to remove particulates before adding an aliquot to the Qubit working solution.
- Inaccurate pipetting: The Qubit assay will accept 1-20 µL of sample, but pipetting very low volumes, especially 1-2 µL is typically very inaccurate, especially with viscous samples. If possible, pipette at least 5 µL for more consistent results.
- The temperature of the assay is changing: Make sure that the Component B buffer is at ambient temperature before use and avoid leaving the samples in the Qubit instrument or near an exhaust fan or other heat source that would warm up the samples.
- Contamination in buffer causing high background: High buffer contamination will show up as an increase in the relative fluorescence (RFU) of the background, measured with the standard tube 1 blank, and eventually will trigger a “Standards Incorrect” error warning on calibration. Replace the kit.
- Sample buffer contains detergents: Qubit protein assay is a detergent-based assay, utilizing an environmentally sensitive dye that fluoresces in the presence of detergents; therefore, only very low concentrations of additional detergents are tolerated in the assay, as listed in Table 2 in the manual (https://tools.thermofisher.com/content/sfs/manuals/Qubit_Protein_Assay_UG.pdf).
- Sample buffer contains other components that are affecting the assay: The Qubit protein assay is generally tolerant of reducing reagents, salts, free nucleotides, amino acids, DNA, and solvents. Table 2 in the manual (https://tools.thermofisher.com/content/sfs/manuals/Qubit_Protein_Assay_UG.pdf) lists acceptable concentrations for many common contaminants. If you have a contaminant you think is affecting the quantitation, then prepare duplicate standard tubes, and spike the contaminant into one set of tubes and run them as samples. If the effect is not too substantial, then spike the buffer into the standards when performing the calibration to account for buffer composition effects.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
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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.
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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).
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
The lowest protein size limit for these reagents has not been determined, although proteins as small as 6000 Da have been accurately quantitated. Quantitation accuracy of small peptides would likely be variable and dependent on the composition of the peptide. We would recommend using the CBQCA Protein Quantitation Kit for quantitation of small peptides. For quantitating peptide digest mixtures for mass spectrometry applications, we recommend using the Quantitative Colorimetric Peptide Assay (Cat. No. 23275) or Quantitative Fluorometric Peptide Assay (Cat. No. 23290).
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We do not recommend that you use the Quant-iT Protein Reagent or Qubit Protein Reagent to quantify proteins in the presence of any detergents, surfactants, lipids or other chemicals that can either displace the dye from a hydrophobic domain, disrupt lipid structure, or add a lipophilic/hydrophobic componentto the solution. The dye is environmentally sensitive; when it binds to hydrophobic pockets/domains, inserts into liposome lipid layers, or is dissolved in organic solvents, its fluorescence output increases relative to its fluorescence in aqueous solutions, potentially providing a higher background. Of course, this assumes that the liposome is not composed of anything that can quench fluorescence.
You may use the dye to quantitate purified protein and possibly a pure liposome sample (assuming that the solution the dye is dispersed in does not disrupt liposome structure), but it would be exceedingly difficult to quantify either as a mixed population.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
All the above assay kits come with either concentrated assay reagent and dilution buffer or a pre-diluted quantitation reagent and protein standards. The EZQ Protein Quantitation Kit also comes with a specially-designed 96-well microplate and filter paper that fits inside this microplate.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
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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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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).
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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
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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.
Find additional tips, troubleshooting help, and resources within our Protein Assays and Analysis Support Center.
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.
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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.
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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.
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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).
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Pierce Bradford Protein Assay Kit and Pierce Bradford Plus Protein Assay Kit are variations on the use of Coomassie G-250 dye as a colorimetric reagent for the detection and quantitation of total protein first reported by Bradford in 1976. The Thermo Scientific 660 nm Protein Assay is a dye-based reagent that offers the same convenience as Coomassie-based assays while overcoming several of their disadvantages. In particular, the 660 nm Assay is compatible with most detergents and produces a more linear response curve (the detailed assay chemistry is proprietary). Our fluorometric protein assays are also based on dye binding chemistries.
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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.
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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.
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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.
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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.
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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.
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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.
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We offer a Quant-iT Protein Assay Kit (Cat. No. Q33210), which is more sensitive than standard absorbance-based assays and can quantitate proteins from 0.25-5 µg. The signal is unaffected by many common contaminants, such as DTT, beta-mercaptoethanol, amino acids, and DNA. Imidazole at a final concentration below 1.25 mM is acceptable. Above that concentration, the imidazole begins to interfere with the assay.
Please note, imidazole does absorb at 280 nm, and the absorbance varies with concentration. So to be perfectly accurate, each eluted fraction should be blanked against its elution buffer.
Find additional tips, troubleshooting help, and resources within our Protein Purification and Isolation Support Center.