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Requirements: complete as required
For Assignment 1, Look at any heading in green for this assignment, the heading is in Green and Says Assignment 1: (If you have trouble seeing green each part also says Assignment 1)
This document will be used as the basis for your final formal paper though additional data and a narrated slide show on the procedures will be provided on the canvas shell as well.
First you will write a Methods, then a Results, then an Introduction, and a Finally a Discussion of this experiment.
Final Formal Paper: Then you will revise the parts of the paper, based on the comments provided by your instructor or the problems noted on the rubrics for each of the individual assignments. You will combine them into a Formal paper that includes an Abstract.
The parts will be assigned after the assignment at the end of this document is graded for accuracy of the calculations involved. Additional Raw data of a similar nature will be provided for the results section. This experiment coincides with the pipetting exercise in lab 4 as it also is using the Bradford assay. The purpose of the experiment is to determine the amount of protein in a sample and compare that to what the labeling on the package says the protein content is for that product.
All cells contain hundreds of different biomolecules, including proteins, carbohydrates, lipids, and nucleic acids. These terms refer to classes of compounds and there are actually many types of proteins, carbohydrates, etc. The total amounts of these different molecules vary from cell to cell or from tissue to tissue. An initial step that is often done to characterize a particular cell type is to determine the total amounts of the different types of biomolecules per cell. This is usually accomplished by extracting the molecules from a collection or set of cells and then by doing a spectrophotometric assay to measure the total amount of a certain type of molecule quantitatively. This involves the same basic spectrophotometric methods you learned in last week’s lab.
Assignment 1: A data set for a protein sample whose protein content is unknown as well as a data set for a standard is be provided.
Standard Curves
When you do not know the molar extinction coefficient (E) for a particular molecule or are uncertain about the linearity of the relationship between concentration and absorbance, you could use spectrophotometry to make quantitative measurements if you first construct a standard curve. A standard curve is a graph that shows the relationship between the amount of a particular compound in a solution and the absorbance of that solution.
Once a standard curve has been created, it can be used to determine the amount of the compound of interest in an unknown solution. The absorbance of this unknown solution is first measured using the same instrument at the same wavelength as the standards. You can use the equation for the linear region of the standard curve to create a conversion factor for relationship between absorbance and amount of unknown in a solution.
Within the linear region of a standard curve, the straight line has the formula:
where m is the slope of the line and b is the Y intercept. If b = 0, that is, the line goes through the origin at 0,0, you can use the slope of the line (m) as the conversion factor since it directly gives the relationship between x and y. You can then divide the absorbance of the unknown sample by the conversion factor to determine the corresponding amount.
Spectrophotometric Assays for Proteins
In this hypothetical experiment, we have used the Bradford Method because it is particularly easy to do. This method was first described by Bradford in 1976. It is based on the binding of the dye Coomassie Blue G-250 in a phosphoric acid solution to proteins (Figure 4.5).

While free Coomassie Blue G-250 has an absorption maximum at 465 nm and solutions of it are brown in color, the dye-protein complex has an absorption maximum at 595 nm and solutions containing these complexes are blue in color (Figure 1). The Bradford method is particularly easy to use and only takes about five minutes.
Any protein can be used to construct a standard curve for the Bradford assay, but the most commonly-used protein is bovine serum albumin (BSA), which is found in serum from cows. BSA is normally used to transport fatty acids through the blood and is commercially available at relatively low cost. It should be noted that the use of the standard curve based on a particular protein is only a matter of convenience. In an extract of a protein shake, there is no BSA! The statement that a particular solution has a protein concentration of 2.58 mg/ml only indicates that the solution contains as much “apparent protein” as a 2.58 mg/ml solution of BSA. The solution is a mixture of many different proteins in varying amounts, which only show as much reaction as the corresponding amount of the standard protein.

Basic Procedure for the Bradford Assay
To do the Bradford protein assay, you will add a series of solutions to a brand new 13 x 100 mm test tube. New tubes are used to avoid any contamination introduced into the tubes from previous experiments. The following steps should be done in the order given:
add water to the tube with a micropipetter (0 to 100 μl)
add the BSA standard or another protein solution to the tube with a micropipetter (5-100 μl)
mix the complete sample (100 μl) by inversion
add 3.0 ml Bradford reagent and mix by inversion
incubate at room temperature for 10 minutes
read absorbance at 595 nm
Example of a Protein Assay (Help with calculations for Assignment 1)
To illustrate how a standard curve is made and used, consider the following example. As part of an enzyme purification procedure, it was necessary to determine the protein concentrations of a series of fractions or samples containing the enzyme of interest. The BCA reagent was used in this case. A protein standard curve was first created by varying volumes of a bovine serum albumin solution to a series of tubes containing a total volume of 2.0 ml. Table 1 shows the absorbance values at 562 nm for different amounts of BSA.
Table 1. Absorbance at 562 for different amounts of BSA

To make a standard curve, the absorbance of each solution was plotted as a function of the amount of protein (Figure 2)
Notice that the amount of protein in μg is plotted on the X axis and the absorbance at 562 nm is plotted on the Y axis. The data points fall on a straight line from 0 to 50 μg of protein. You can use the line to create a conversion factor relating absorbance to amount. Since 10 μg of protein meets the line at an absorbance value of 0.1, the conversion factor is:
Suppose now that 10 μl of one of the fractions gives an absorbance of 0.251 under the same conditions. What is the protein concentration in mg/ml?
You can answer this question either by reading the numbers off of the graph or by using the conversion factor.

From the graph, 0.251 corresponds to about 26 μg.

From the conversion factor,
Remember that 1 μg/μl is the same as 1 mg/ml, so you don’t really need to multiply through by all of the metric conversion factors that this shows! Also remember that if the sample is diluted you must adjust for dilution.
Also note that you cannot use the absorbance of if it is beyond the range of the standard curve meaning that your curve does not contain the point. Even though the line appears to be linear, you have no way of knowing that it continues indefinitely. In fact, most standard curves will deviate from linearity at high absorbance values or high amounts.
Background for Spectrophotometry:
Quantitation of Absorbance
The total amount of light absorbed at any particular wavelength (the absorbance) is determined by three factors: 1) the absorption characteristics of the molecules of interest; 2) the pathlength or distance through which the light must travel; and 3) the concentration of the absorbing molecule. The absorption of light is usually measured in an aqueous solution, where the intensity of the light passing through the sample decreases exponentially with the thickness of the water layer. The intensity of the light passing though the sample also decreases exponentially with the concentration of the solute. These factors are summarized in the following expression, which is called the Beer-Lambert law.

The Beer-Lambert Law can be calculated using the expression:
A = log10 Io = E c l
I

where A is the absorbance of the solution, Io is the intensity of the incident light, and I is the intensity of the transmitted light; E is the molar extinction coefficient; c is the concentration of the absorbing solute; and l is the pathlength of the light.

The concentration of the solute (c) is usually expressed in moles/liter (M) and the pathlength of light (l) is expressed in cm. The molar extinction coefficient (E) is an intrinsic characteristic of each molecule at a particular wavelength. It is numerically defined as the absorbance of a 1.0 M solution of the molecule of interest in a 1.0 cm light path. Because E has the units of liter cm-1 mole-1, absorbance itself is a parameter with no units. The larger the value of E, the more a compound absorbs at a particular wavelength.

Because molecules vary in structure and have different electronic states, they have characteristic absorption or emission spectra and unique extinction coefficients. Some biologically-important compounds absorb light in the visible region of the spectrum (400-700 nm), some in the ultraviolet (UV) region of the spectrum (200-400 nm), and some in the infrared (IR) region of the spectrum (700-1000 nm). Fluorescence usually occurs in the visible or infrared region. Biochemical assays for biomolecules are usually based on absorption in the UV or visible region and fluorescence in the visible region.

Quantitative assays based on the absorption or fluorescence of light are usually performed using a spectrophotometer or spectrofluorometer. These instruments usually compare the absorption or fluorescence of a solution containing the compound of interest (the experimental sample) to one that does not contain that compound (the reference sample or blank) (Figure 3). The difference in amount of light absorbed or emitted can be expressed as an absorbance value, a percentage of the incident light transmitted, or a relative fluorescence.
Figure 3. The major components of a spectrophotometer.

Light is produced from a lamp, which usually has a tungsten filament bulb for light in the visible region and a deuterium discharge bulb for light in the UV region. The light then enters a monochrometer, which splits the light into different wavelengths using a prism or diffraction grating. Light of a selected wavelength then passes into the sample compartment. Some spectrophotometers are single-beam instruments which have only one light path. The reference and experimental samples are compared by moving first one into the light path and then the other. Other spectrophotometers are double-beam instruments in which the monochromatic light is split into two beams that pass simultaneously through the reference and experimental samples. Light transmitted by a sample then enters a detector, which is usually a photomultiplier tube that converts light energy into an electric current. The current coming from the experimental sample is compared with that coming from the reference sample and the result displayed on a digital or analog meter, a cathode ray tube, or a recorder. Fluorescence measurements are made in a similar way. A spectrofluorometer has the same basic components as a spectrophotometer, but it also has a second monochrometer to select the wavelengths of emitted light that are detected and used in the measurements. In most spectrofluorometers, the detector is positioned at a 90 degree angle from the sample to avoid the effects of normal transmitted light. Many different types of spectrophotometers and spectrofluorometers are commercially available.

The instrument should be used in the following way (Figure 4):

Be sure the power cord is plugged into a grounded 120 Volt outlet.

Turn on the power switch on the back of the instrument. The instrument will go through a short power-up sequence that takes about 2 minutes. Then allow the instrument to warm up for 15 minutes before taking any readings.

Press the A/T/C button on the key pad to select absorbance.

Press the nm(UP) or nm(DOWN) buttons on the key pad to select the wavelength to be used.

Lift up the cover of the sample compartment and insert a 13 x 100 mm test tube containing the reference solution. Be sure to wipe the outside of the tube first with a Kim-Wipe to remove any liquid or fingerprints. Be sure to insert the tube correctly so that light passes through the clear walls. Close the cover of the sample compartment.

Press the 0 ABS/100% T button on the key pad to set the instrument to 0 absorbance. The zero reading will appear on the LCD display.
This experiment has several parts but they must be done sequentially.
The following is a flow chart of the information generally used for this experiment.

The following information is provided for you so you can write a Methods section on this experiment you can assume that the experiment was done exactly like it is described in this document:
Assignment 1: All data from the experiment is be provided to you by your instructor at the end of this document.
Preparation of a Food Source Extract
The purpose of this part of the experiment is to prepare an extract of your food source in a simple buffer solution so that its protein content can be determined.
Look closely at the nutritional label on the side of the package of food you brought to the lab. Note the serving size and the number of grams of protein per serving.
Open the package and measure out one-tenth serving size. Depending on the product, it might have a certain weight in grams or ounces (remember 1.0 g = 0.0353 ounces) or a certain volume in liters or cups (one 8 ounce cup = 250 ml). Balances and measuring materials will be available for you to use. Smaller amounts can be used if it looks like there is a very large amount of material. Consider that the total volume of liquid you will add is 20 mL
Transfer the material to a clean beaker.
Add 50 mL of 0.1 M potassium phosphate buffer, pH 7.0 to the food material.
Stir the material for two minutes or until no clumps remain if using a protein powder.
Decant the suspension into a graduated cylinder and measure the volume in milliliters.
Then transfer the suspension to a clean flask and save it for the protein assay in Section C. This is your protein extract.
Record the following information on your food source:
What was the food source that you used?
________________________________
What was the designated serving size? What part of a serving did you use?
_______________________________ _________________________________
What was the labeled protein content in grams/serving?
_______________________________
How many ml of extract did you obtain after homogenizing one or part of one serving? If you used less that an entire serving note that also.
________________________________
Setting up a Protein Standard Curve
The purpose of this part of the experiment is to prepare a protein standard curve, using bovine serum albumin (BSA) as the protein standard and the Bradford Reagent. You will set up a series of tubes with varying amounts of BSA and a constant amount of Bradford reagent. By plotting the number of micrograms (μg) of BSA on the X-axis and the corrected absorbance on the Y-axis, you will be able to generate a standard curve that can then be used to determine the protein concentrations of your unknown solution.
At the beginning of the lab, turn on the Genesys 20 spectrophotometer and allow it to warm up for 15 minutes. Set the wavelength to 595 nm.
You will be provided with a 0.5 mg/ml stock solution of bovine serum albumin (BSA). Remember that 0.5 mg/ml is the same concentration as 0.5 μg/μl.
Set up 17 13 x 100 mm glass tubes as shown in the following table. Notice that you will be testing volumes of BSA from 10 to 100 μl in duplicate. Notice also that BSA will be added to the water to give a total sample volume of 100 μl. 3.0 ml of the Bradford reagent then will be added to each tube. (THE ORDER IN WHICH YOU ADD THESE IS IMPORTANT!!)
Using micropipetters, add the water to the tubes first. Then add the BSA solution. It will help the accuracy if you use a new tip for each sample. Flick the tubes with fingers gently to mix.
When all of the samples have been prepared, add 3.0 ml of Bradford Reagent to each tube using a Repipetter. The instructor will demonstrate how to use this device.
Cover each tube with part of a square of Parafilm and invert it several times. This is better than vortexing the samples because it does not generate a lot of foam.
Allow the tubes to sit at room temperature for 10 minutes.
Use the solution in tube # 1 to set the instrument to zero absorbance since this “blank” contains only water and Bradford Reagent.
Measure the absorbance of each tube at 595 nm using the chart shown below. The tubes will fit directly into the cuvette holder of the Genesys 20 spectrophotometers.

Enter your raw data for the absorbance measurements of the samples from the bovine serum albumin (BSA) standard curve
Table ____ Title: ____________________________________________
Calculate the average absorbance value for each of the duplicate samples
Then determine the total amount of protein in each of the pairs of tubes. Remember that the stock solution is 0.5 mg/mL or 0.5 μg/μL. Example: if tube 2 contains 5 μL of solution you would conclude that it also contains 2.5 μg of protein since it contains 0.5 μg/μL of BSA.
Using a piece of linear graph paper, plot the average absorbance values as a function of the amount of BSA in each pair of tubes. Draw a “best fit” straight line through data points with a ruler. This line should go through the origin since 0 BSA = 0 Absorbance. The line should pass through or come close to most of the data points. You might find, however, that the standard curve becomes nonlinear at high protein concentrations. (Insert a copy of this graph into your lab manual after this page) Note: non-linearity is not slight deviations; instead it is the beginning of a slight curve and should not appear in these data since you are using fairly small concentrations.
Discuss the graph with the instructor. If it looks good, you can proceed to the next part of the experiment. If some of the points deviate badly from the straight line, set up new tubes for those amounts of protein and repeat the assay.
Once you get a good standard curve, make up a conversion factor relating the absorbance at 595 nm to the amount of protein (______ A595/μg). This conversion factor is the absorbance of the sample, divided by the the slope of the line in the linear region of the standard curve and can be calculated from any convenient set of points within the linear region.
Determination of Protein Concentration of the Food Source Extract
The purpose of this part of the experiment is to determine the protein concentration of your food source extract. Since you do not know the concentration of the unknown solution, you will need to make several dilutions so that some of your protein samples will fall within the range of the standard curve.
Make 3 serial 1/10 dilutions of your unknown solution in the following way. Add 900 μL of pH 7.0 phosphate buffer to each of three 1.5 ml microcentrifuge tubes. Mix the extract you prepared earlier and then and add 100 μL of it to the first tube. Close the cap and invert several times to mix. Then add 100 μL of the 1/10 dilution to the second tube to make a 1/100 dilution. Again, close the cap of the second tube and invert to mix. Finally add 100 μL of the 1/100 dilution to the third tube to make a 1/1000 dilution. Close the cap and invert to mix. It may be a good idea to vortex these before proceeding to the next step. Then using these diluted solutions set up an assay as described in #2 below.
Set up a new protein assay as shown in the following table.
Table ____ Volumes of liquid used to set up an assay of ______________protein extract.
Note that by following this protocol, you would be testing 10 μL, 30 μL, and 70 μL volumes of each of the dilutions. Again, the total volume in each tube before adding the Bradford Reagent will be 100 μL.
Using micropipetters, add the water to the tubes first. Then add the extract you just made whose protein content is theoretically unknown. Again, it will help the accuracy if you use a new tip for each sample.
When all of the samples have been prepared, add 3.0 mL of Bradford Reagent to each tube using a Repipetter provided.
Cover each tube with part of a square of Parafilm and invert several times. This is better than vortexing the samples because it does not generate a lot of foam.
Allow the tubes to sit at room temperature for 10 minutes.
Measure the absorbance of the solution in each tube at 595 nm and record the value. You will probably find that some of the solutions are very dark and give absorbance values beyond the range of the standard curve. You may also find that some of the solutions are very light and give absorbance values that are too low (<0.05) to be meaningful or very accurate.
Enter the raw data for the unknown protein sample here
Table _____ Raw data giving absorbance of ___________________ prior to any calculation of protein content.
Assignment 1: this part can be use for help with the calculations below in Assignment 1.
For analysis of protein content use only those samples whose absorbances fell within the linear range of your BSA standard curve these samples will fit into the USABLE category. You can either interpolate directly along the line of the standard curve or use the simple conversion factor (slope of the line) derived from it. Calculate the amount of protein in μg in each of the usable sample. (Absorbance measured from your unknown protein) = Slope of the line (from your graph) multiplied by (x).+ 0 since our y intercept in this case is zero. See the introduction for more specifics on how to do this calculation.
Enter the amount of protein in each USABLE sample in the following table:
**Enter N/A for samples that are not usable note that you do not include any of these values in your paper these would be included only in the assignment.
Then, correct for the volume used in each sample and the dilution factor to calculate the protein concentration of the original protein suspension in mg/mL. For example, suppose that 30 μl of a 1/10 dilution turns out to contain 13.7 μg of protein. The protein concentration is then:
Include the calculations in your lab notebook.
If you have several samples that give absorbance values within the range of the standards, calculate the protein concentration for each sample separately. Then average the values to get a single protein concentration for the original solution. Record that value here or in your lab notebook.
Final Average protein concentration in _______________________
(food tested)
________________________mg/mL
Final Calculations
The purpose of this part of the experiment is to complete the calculations necessary to determine if the nutritional label on your animal or plant food source is accurate.
After completing Section C, you should have a value for the protein concentration of your food source extract in mg/ml. Multiply this value by the total volume of the extract that you measured in Section A to get the total amount of protein from your sample.
For example, if the protein concentration was 9.42 mg/ml and the total volume of the extract was 53.5 ml, the total amount of protein can be calculated as:
All of this protein came from the initial amount of material you added to the liquid in the blender as measured in weight or volume. Depending on whether you used a full serving size, one-half, or one-fourth of a serving size, calculate the total amount of protein per serving size. Compare the value from the data provided with the value on the nutritional label. Record your final values and include a brief comment regarding your protein source and how it relates to the amount of protein you expected to find.
Expected (from label): ___________________g protein/serving
Observed (from your calculations): ___________________ g protein/serving
The data below is to be used for Assignment 1 additional data will be provided for the Results section:
Assignment 1 Starts here:
Absorbance readings from BSA Standard: Assume a 0.5 mg/ml BSA standard was used.
Table 1 Title: BSA standard readings at 595 nm based on vol of BSA.
(10 points). Complete the table above (3pts) and Create a graph (5pts) of the BSA standard curve from the data provided (Table 1). Draw the best fit straight line that is possible through most of the points. The line should go through the origin (0,0) since all of the samples were read against a “blank” containing 0 protein that was set to 0.000 A.
Define a conversion factor relating absorbance to the amount of protein. This is the same as the slope of the line from your graph.
y. = mx + b, where m = slope of the best fit line and x = mg of protein, b = y intercept which in your case should be set to 0.
Absorbance is on the y axis of your graph and mg of protein is on the x axis
(2pts) Your conversion factor relating the absorbance at 595 nm to the amount of protein
(______ A595/μg) derived from your graph. This conversion factor is the absorbance of the sample, divided by the slope of the line in the linear region of the standard curve and can be calculated from any convenient set of points within the linear region.
Absorbance readings from dilution of samples:
These are the raw data you will use for this problem:
2. (10 points) Calculate the Protein Concentration of the Food Source Extract.
For this use your standard curve from which you have calculated a conversion factor relating the absorbance at 595 nm to the amount of protein (______ A595/μg). This conversion factor is the absorbance of the sample, divided by the slope of the line in the linear region of the standard curve. Make sure to avoid using any samples whose absorbances are above the range of your standard curve. It is also important to avoid using samples with values below .01 because the last digit on the spectrophotometers used is variable and may give a value even if the value is 0. (hint: there are five) put this information in the table above
Then for each sample above, correct for the volume used in each sample and the dilution factor to calculate the protein concentration of the original protein suspension in mg/mL. For example, suppose that we use a 30 μl sample of a 1/10 dilution. The protein concentration (from the left column of the table above would then be:
Include the calculations here for each sample usable sample. (5 pts) (Again there are five)

After you know the mg/ml of protein in the tested samples calculate their average.
Figure _. Label of Breakfast Essentials Nutritional Drink.
3 (10 points) . After completing the calculations above you should have a value for the protein concentration of your food source extract in mg/ml. Multiply this value by the total volume of the extract that is provided above to get the total amount of protein from your sample.
For example, if the protein concentration was 9.42 mg/ml and the total volume of the extract was 53.5 ml, the total amount of protein can be calculated as:
Food source used. (See provided label) (1pt)________________________________
What was the designated serving size (See provided label).(1pt) _____________
Amount of a serving used ____1/10th____________
labeled protein content in grams/serving (See provided label). (1pt)______________
Number of ml of extract obtained after homogenizing one part of one serving. ____55.2 mL_____
Then multiply but the amount of a serving used. In our case 1/10th so to get back to an entire serving multiply by 10. (2pts)
All of this protein came from the initial amount of material added to the liquid in the blender as measured in weight or volume. Compare the value from the data provided with the value on the nutritional label. Record your final values and include a brief comment regarding your protein source and how it relates to the amount of protein that was expected.
Expected (from label): ___________________(1pt) g protein/serving
Observed (from your calculations): ___________________ (2pts) g protein/serving
Compare your value for the protein content with that shown in the nutritional label? How can you explain any differences between these values? (2pts)

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