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Lab #1: Determining Sample Protein Concentrations
In this exercise, you will learn how to generate and use a standard curve to determine protein concentrations from an unknown sample. This is often an important first step when analyzing protein samples, for example from a cell lysate. Here, you will gain practice with micro pipetting, colorimetric assays and spectrophotometry, serial vs parallel dilutions, generation of a standard curve, and calculation of protein concentrations. Gloves must be worn at all times while performing experiments in the lab, and goggles worn here when pipetting and vortexing samples containing Coomassie dye. Glass tubes are disposed of in glass waste containers. Upon completion of this lab exercise, you should be able to:Describe how Coomassie brilliant blue G-250 dye can be used to detect proteins in solutionDetermine how a set of known protein sample concentrations can be utilized to determine the concentration of an unknown sample through the use of a Coomassie colorimetric assayIdentify the unknown sample dilution(s) that allows you to most confidently determine the protein concentration of your unknownCalculate the unknown sample’s concentration taking into consideration dilution factors, where neededIdentify potential sources of variation in the experimental procedure and establish techniques to help minimize experimental errorSuggest when to use serial vs parallel dilutions and why each can be beneficial depending on the particular experimental conditions Background You will be using a Coomassie Plus reagent(Thermo Scientific) to quantify protein by spectrophotometry, a procedure similar to the original Bradford assay (Bradford, 1976) for protein quantification. The Coomassie reagent includes a dye, Coomassie brilliant blue G-250, which produces a color change in the solution upon binding to protein. The dye exists in three different charged forms, with equilibriums of these free dye forms driven by the pH of the environment. Positive, neutral, and negative charges result in red, green, and blue forms of the dye, respectively. In an acidic environment (low pH), like that of the Coomassie Plus reagent, most of the dye will exist in the red, positive charged (doubly protonated) form due to the excess of H+ions in the solution. However, studies have suggested that it is the blue, negative charged(unprotonated)form of the dye that binds protein (Chial and Splittgerber,1993). The blue form of the dye binds to protein through non-covalent interactions with arginine and other basic residues, and Van der Waals and hydrophobic interactions with aromatic amino acid side chains are also thought to be involved (Compton and Jones, 1985).So based on this information, how do we understand the assay to work? In the acidic environment of the Coomassie Plus reagent, a small percentage of dye exists in the blue form. In the presence of protein, the blue form will bind, generating blue dye-protein complexes. This binding decreases the 2amount of free blue dye in solution, shifting the equilibrium to restore blue dye levels, providing more blue form of the dye available to bind protein. The red form absorbs maximally at a wavelength of 470nm and the blue form absorbs maximally at 595nm(Chial et al, 1993). Therefore, the more protein, the more blue dye-protein complexes formed, the more blue color and the higher the absorbance at 595nm by spectrophotometry. Why use an acidic environment for this assay? A low pH, in the absence of protein, results in predominantly red form of the dye, which will provide a nearly “negative” reading at 595nm. The amount of blue dye in an acidic environment is proportional to the amount of protein. Absorbance readings at 595nm will therefore serve to reflect protein concentrations in the solution.(The green form is also present, but has a maximum absorbance at 650nm, so does not interfere significantly with measurements at 595nm.)Alternatively, you could measure proteins directly by spectrophotometry. Proteins absorb maximally in the UV range at a wavelength of 280nm. However, other biomolecules that may be present in solution can also absorbUV light, complicating interpretationof your absorbance reading. For example, DNA absorbs UV light at 280nm (maximally at 260nm),as does RNA. The Coomassie colormetric assay can therefore oftenallow for more specific analysis of proteins.ReferencesBradford, M.M. 1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem, 72:248-54.Chial, H.J and Splittgerber, A. G. 1993. A comparison of the binding of Coomassie brilliant blue to proteins at low and neutral pH. Anal Biochem, 213: 362-69.Chial, H.J., Thompson, H.B., and Splittgerber, A.G. 1993. A spectral study of the charge forms of Coomassie blue G. Anal Biochem, 209:258-66.Compton, S.J. and Jones, C.G. 1985. Mechanism of dye response and interference in the Bradford protein assay. Anal Biochem, 151:369-74.3Your objective: Use a Coomassie colorimetric assay to determine the concentration of your unknown protein sampleStep 1: Generate a standard curve from known concentrations of bovine serum albumin (BSA)Step 2: Use your standard curve to determine the protein concentration of your unknownStep 3: Assess areas for improvement/optimization for subsequent protein quantificationin the labReagents/Materials/Equipment available to you:CoomassiePlus Reagent (Thermo Scientific)Visible light Spectrophotometer2mg/mL BSA(~150ul)Deionized (DI) H20Pipetmen and pipettesUnknown protein sample (~100ul)Glass tubesfor spectrophotometry1.5mL tubesfor making standards and sample dilutionsVortexGuidelines for generating your standard curve:By establishing a standard curve, you will then be able to place the absorbance value for your unknown on this curve to determine its concentration. Here, you will first generate 4newsamples of BSA in 1.5mL tubes, at set concentrations. These, along with the 2000ug/mL stock,will give you 5 different BSA concentrations for your standard curve:Based on a known linear working range of BSA concentrations in this assay, set your lowest standard concentration to 100ug/mL and the highest to 2000ug/mL (which is the concentration of your stock BSA solution). You can then select 3 concentrations in between those values to utilize to generate your standard curve. Use the tables below to establish your conditions.You are also asked to calculatethe total ugof BSA in 10uL here because this is the volume of each sample that will be added to the glass tubes for measurement (see general procedure details below).Be careful with unit conversions!Pay attention to ug vs mg and uL vs mL here!StandardConcentration (ug/mL)Total ug BSA in 10uL110023452000Based on the above concentrations you have selected to use for your standards, now use the following table to determine the volumes needed to generate these from the 2mg/mL BSA stock provided. You will make a total of 40uL of each standard. This is sufficient volume to perform the experiment and will provide some extra in case you want to run any replicas or need to repeat a measurement.You do not need to generate the 2000ug/mL standard because this is the same concentration as the BSA stock 4(2mg/mL) you already have.The BSA standards should be diluted in deionized (DI) H2Oin 1.5mL tubes.StandardConcentration (ug/mL)Total volume to makeVolume of 2mg/mL BSA stock (uL)Volume of DI H20 (uL) 110040uL240uL340uL440uL52000n/an/an/aOnce your TA/Instructor has checked your work and demonstrated how to utilize the spectrophotometer:Generate your standards in new, labeled 1.5mL tubes.Be sure to label each tube on the cap and/or on the side of the tube before you get started.Perform the Coomassie assay togenerate your standard curve (see additional guidelines below for experimental procedure).Be sure to record all steps, in the order youperformedthem,in your lab notebook.Ask your TA/Instructor ifyou are not certain about the level of detail you need to include.General ProceduresCoomassie Assay:This procedure is utilized for generating the standard curve and for analyzing your unknown samples. You willestablish your curve first before you start working with your unknown sample.1.Add 1mL DI water to each glass tube2.Add 10uL of standard or unknown sample (10uL DI water for blank)3.Add 1mL CoomassiePlus Reagent(always add the Coomassie dye last)4.Gently vortex and incubate at room temperature (RT) for 3-5min. Try to be consistent with the time interval, from addition of Coomassie reagent to measurement of absorbance in the spectrophotometer,for each sample.5.Blank the spec prior to recording measurements for samples: Set the spectrophotometer to 595nm. Place the tube containing 1mL DI H20 + 10uL DI H20 + 1mL Coomassie in the spec and zero the instrument.You only need to blank the spec when you start, not between each sample.Alternatively, you can blank the spec with water only (no dye)and then subtract out the measurement of your water only + dye control from other samples. Your TA/Instructor will provide guidance for which method to use here.6.Proceed with measurements of samples, recording absorbance readings at 595nm.Once you have completed these two tables, ask your TA/Instructor to check your experimental design and calculationsbefore you proceed.5Note: It is bestto use the same solvent to blank and measure standards as is used in your unknown sample. Here that is DI water, but be aware that often the protein sample you will need to quantify is in a particular buffer (perhaps different solvents and often solutes are present in the sample as well) depending on how it was isolated/purified. For example, crude protein samples from cell extracts are oftenextractedin lysis buffer. Components in the buffer can often impact light absorption by spectrophotometry, making it inaccurate to compare absorption of proteins fromdifferent buffer/solution sources.Graphing your standard curve in excel:1.Enterstandard sample concentrations (ug/mL) in one column and corresponding absorbance values in an adjacent columnin an excel spreadsheet.2.Highlight both columns and select insert graph.3.Choose scatter dot plot format with concentration on the x-axis and absorbance on the y-axis.4.Add a linear trendline and R2value to the graph.Depending on the version of excel, these options may be available under the Design tab and Add Chart Element or through Format Chart Area, Chart Options. To show the line equation and R2value on the graph, select More trendline options.Analysis of your Standard CurveThe standard curve can plot the absorbance vs the concentration (e.g. ug/mL) or the absorbance vs the amount (e.g. ug). Here, because we are always adding 10uL of each standard and unknown sample to the glass tube, with the same volumesof water and dye, we can simplify our analysis and follow the concentration of the sample examined. However, if you chose to, you could also calculate and plot the total ug of protein added to the glass tube and then calculate back to determine the concentration of the sample added.Protein standard curves are not completelylinear in nature when plotting absorbance vs concentration/amount.There is a range of protein concentrations whereabsorbance is directly proportional to concentration. However, absorbance readings falling outside of this linear working range do not provide accuratemeasurementsof protein concentration. Too little protein can fall below the threshold of detection,and too much protein can saturate the reaction. Here, you have guidance for which concentrations to use at each end of your standard curve range. These were designed in efforts to allow you to work within the known linear range of BSA in this assay and to span as much of that range as possible.Once you have measured and plotted the absorbance values for your standards to generate your standard curve, discuss with your TA/Instructor to determine if you are ready to proceed with measurement of your unknown sample. Does your graph look linear within the range tested? How confident are you in your measurements? What does the trendline equation tell you? R2value?6Analysis of your Unknown Protein SampleNote which unknown sample your group is analyzing: _______Typically you would set up your unknown sample tubes along with your standards to keep all conditions in the Coomassie assay as consistentas possible, processing them together. However, because this is the first time we are constructing a standard curve, here you will generate the curve first and then analyze your unknown samples once you have generated a functional standard curve. Take a look at your standard curveonce you have it completed. Ideally where would you like absorbance of your unknown sample to fall within the curve to give you most confidence in accuracy of your measurement? What happens if your unknown protein sample falls below or above the linear range of your standard curve? If your unknown falls below the linear region of your curve, there is not much we can do using this particular assaywith your current sample. If your unknown falls above the linear range of your curve, that issue can be easily resolved!Dilutethe sample to a dilution that now falls within the linear range. Then the dilution factor can be used to calculate the concentration of your original (undiluted) unknown.Typically, researchers will perform several 3-10 fold serial dilutions of their unknown sample if the original sample concentration is too high whenmeasureddirectly.How to determine if dilutions are needed and if so, how to estimatethedilution(s)needed other than through trial and error? Hint: Take a look at the color of the sample whenadding Coomassie reagent to the glass tube with your10uL ofunknown. You should be able to approximate by finding dilutions that fit within the color range of your standards.Keepin mind there is only ~100uL in the unknown stockyou were provided with,so be careful to plan accordingly.If you use serial dilution to make your unknown dilutions, be sure to calculate how much total volume you need of each dilution. For example, if you made three 5 fold serial dilutionsfrom your original 1:1 (100%) sample stock:1:5, 1:25, and 1:125, start by determining how much volume you need of the most dilute, then calculate back from there. Hint: make the math as simple as possible without being wasteful of reagent!Note: Don’t dilute your unknown directly in the original sampletube. Your protein sampleswill often be used in further analysis, with the Coomassie assay serving as simply a means to quantify protein concentration for further applications. Set up new, labeled1.5mL tubes to generate 3 dilutionsof your unknownif you find your sample is too concentrated to analyze directly. Use the table below to guide you, and check your work with your TA/Instructor before proceeding to dilute your unknown.Unknown DilutionsTotal volume to make (uL)Unknown dilution to use as “stock”Volume of unknown“stock”(uL)Volume of DI H20 (uL) 1:1*n/an/an/an/a1:1*Here, notation of dilution denotes part:whole7Use this page to record your absorbance readings for your standard and unknown samplesTube #SampleAbsorbance at 595nmTroubleshooting:Check that your calculations, units and conversionsBe sure to fully mix (by vortexing or pipetting)any dilutions made prior to analyzing themCheck that pipettes are set to correct volumes. Remember these general pipetting procedures:
Be sure to use the correct pipet size for the pipetman in use.
To avoid contamination of samples and reagents, use a fresh tip for each pipetting action, ensure the pipet tip does not come in contact with any other surface, and keep the pipetman upright while in use.
Be sure not to go beyond the first stop when preparing to draw liquid into the pipette.
Keep an eye on what you are doing –pipette slowly and carefully, watching the liquid as it is drawn into the pipet to ensure the pipetman is set properly and watching as the liquid is dispensed to ensure all liquid is transferred.Avoid bubbles.Place the pipet tip right below the surface of the sample as submerging it too far will draw up more sample than intended and can risk contamination if the pipetman itself comes in contact with the sample or inside of the container.It is good practice when adding multiple samples to a new tube to transfer the largest volume first.
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