Laboratory 4: Cell Fractionation - Winona



Laboratory 6 Week 1: Cell Fractionation

Introduction:

Eukaryotic cells are complex and contain many kinds of membrane organelles. For instance, they contain nuclei, mitochondria, vacuoles etc. Two methods exist to study the organelles in more detail. The first is by using a variety of techniques to visualize the nuclei while still inside the cells by microscopy (i.e. cell staining and immunofluorescence). The second method involves suspending the cells in solution, and breaking them open (lysing the cells). Then the various organelles are then separated from each other by centrifugation, which then allows them to be used for further study. It is this second method that we will use in this laboratory.

A procedure called cell fractionation is used to break open the cells and separate the various organelles. To perform cell fractionation, we first will suspend our cells in solution, and then we break open the cells, or lyse them. This will release the organelles inside into solution. Next, we can separate the organelles by centrifuging our solution. By using centrifugation, we can easily separate the various organelles, since the various organelles are of different mass, and density (for instance, nuclei are significantly heavier than mitochondria etc.). During centrifugation, different organelles will pellet at the bottom at specific speeds based on the mass and densities of the organelles. For instance, the heavier (larger) the organelles, the less velocity is needed to pellet the organelle. The lighter (smaller) the organelle, centrifugation must occur at a greater velocity to pellet the organelle.

Therefore, if we want to separate nuclei from mitochondria, we will centrifuge at low speed. At low speed, the nuclei will pellet at the bottom, while the mitochondria will stay suspended in the solution. The left over solution after centrifugation is called the supernatant, and will contain lighter organelles (i.e. mitochondria and dissolved proteins). If we then want to separate the mitochondria from the rest of the supernatant, we can centrifuge at the appropriate speed that would pellet the mitochondria, and then remove the resulting supernatant.

If we first centrifuged or cell suspension at the speed appropriate to pellet mitochondria, we would bring the mitochondria to the bottom of the tube. However, we would pellet everything that is heavier than the mitochondria (i.e. nuclei etc.). Therefore, in order to get nuclei separated from mitochondria, we must centrifuge at the lower speed first to obtain the nuclei, and centrifuge the supernatant at the higher speed to collect the mitochondria. This type of separation protocol is called differential centrifugation. In this procedure, it is possible to separate the organelles to purification because objects a similar size to our desired organelles will also pellet at the same speeds. However, these objects are significantly less concentrated in our suspension. Therefore, our pellets will are enriched for the organelle we are attempting to isolate.

Each pellet that is isolated can then be resuspended by adding solution, and can be called a fraction (for instance, the resuspended nuclear pellet is considered the nuclear fraction). Additionally, a sample of the original suspension is considered a fraction, and is called the crude fraction, as it contains all the organelles and soluble proteins. Lastly, a sample of the final supernatant is also considered a fraction and is called the soluble fraction, and contains everything that was not pelleted by centrifugation.

Using a Centrifuge

There are various types of centrifuges one can use. Various kinds of centrifuges are available: clinical centrifuges seldom run faster than 5,000 rpm but are able to readily sediment cells, nuclei or chloroplasts. High-speed centrifuges go as fast as 25,000 rpm and can sediment smaller organelles such as mitochondria. Ultracentrifuges can produce speeds of 60,000 - 75,000 rpm and can sediment membranous organelles such as microsomes and golgi components. The high speed and ultracentrifuges are always refrigerated and often operated in a vacuum to reduce the heat generated when a rotor is rapidly spinning in air.

To centrifuge your sample, one must first get the appropriate rotor and place it on the pin in the centrifuge. The rotor you choose to use must fit the tubes in which you are placing your cell suspensions. The pin is rotated by the centrifuge motor, thus spinning your rotor. The relative centrifugal force your cell suspension is subjected to is dependent on the speed you are spinning your rotor, as well as the radius of the rotor. You can think of the rotor radius as the distance your centrifuge tube is from the axis of rotation (the pin). In general, your centrifuge tubes are not placed in the rotor in a straight up and down fashion, but are placed at an angle, with the bottom of the centrifuge tube being further from the axis of rotation than the top. The force that that the organelles are being subjected to in order to pellet them is specified by the formula R.C.F. = 1.119x 10-5 (rpm)2r, where RCF is relative centrifugal force, rpm is the revolutions per minute of the rotor and r is the distance (in cm) of the particle from the axis of rotation. Given that the rotor radius is larger at the bottom of the rotor than the radius at the top of the rotor, the RCF will be different along the length of the tube. Can you determine whether the top or bottom of your centrifuge tube will be subjected to greater force? The radius used for our purposes is generally called r(average) and is usually the mean of the maximum and minimum possible radii. Note, many centrifuge rotors come with conversion charts between RCF (which is noted by the units in multiples of gravity (x g)) and rpm.

The laboratory

In the cell fractionation laboratory, we will use differential centrifugation to create fractions enriched for specific organelles. We will first suspend whole plant cells in a mild salt solution (mannitol grinding medium) and lyse them using a mortar and pestle. A sample of the resulting solution will be the crude fraction. By doing differential centrifugation, we will obtain nuclear, mitochondrial and soluble fractions as well. The soluble fraction will contain objects such as soluble proteins, ribosomes and nucleic acids.

Materials

1. Cauliflower or Spinach

2. Razor blades (7)

3. Balances

4. Weighing Dishes (located next to balances in both 284 and 286)

5. Mortar and Pestles (In Cold Room – Chilled)

6. Grinding Sand (Next To Balance in 286)

7. 50 mL Graduated Cylinders (24)

8. Paring knives (2)

9. Ice buckets (6)

10. Oak Ridge Tubes (12) – Chilled in Cold Room

11. Cheesecloth

12. Scissors (3 pairs) – near cheesecloth

13. 250 mL beakers (12)

14. Microfuge Tubes

15. Small glass stirring rods

16. Microscope slides

17. Microscopes

18. Coverslips

19. Methyl Green Pyronin

20. Spectrophotometers

21. 5 mL Microcuvettes

22. Ziplock Bags

23. 13 X 100 test tubes (40)

24. Distilled water (4 x 100 mL bottles)

25. Sharpies (8)

26. Bin for dirty dishes

27. Bin for dirty cuvettes

28. Protein solution (0.1 mg/ml)

Make 2 bottles/100 mL

For 200 mls, take a 400 mL beaker

Fill Beaker with 100 mL deionized water

Weigh out 1 g of BSA and mix into the water

Bring volume up to 200 mL and aliquot into 100 mL bottles

29. Mannitol Grinding Medium

Use a 1 L flask and add 850 mL Deionized water

Add 54.66g D-Mannitol

Add 0.82g KH2PO4

Add 2.42g K2HPO4

pH to 7.2 (This step is extremely important-BE ACCURATE!)

Bring volume up to 1L with deionized water and aliquot in 500 mL bottles (2)

Store Media in the Cold Room

30. Mannitol Assay Medium

Use a 600 ml beaker and add 400 mL Deionized water

Add 27.33 g D-Mannitol

Add 0.41 g KH2PO4

Add 1.21 g K2HPO4

Add 0.38 g KCl

Add 0.51 g MgCl2 X 6 H2O

pH to 7.2 (This step is extremely important – BE ACCURATE!)

Bring volume up to 500 mL with deionized water and aliquot into 2 250 mL Bottles

Store media in the Cold Room

31. 0.4 M Perchloric Acid

Take 34.5 mL concentrated Perchloric Acid

Bring volume up to 1 L with deionized water

32. Protein Dye solution

Obtain a 1 L beaker

Place 50 mL of 95% ethanol in the beaker

Add 600 mg of Brilliant Blue G

Add 750 mL Perchloric Acid and Mix

Bring volume up to 1L with deionized water

Experimental Protocol

Part A: Fractionation of Cauliflower or Spinach Cells (1 head of cauliflower or bag or spinach per lab section)

1. Using a single-edge razor blade, remove a total of 20 g of the outer 2-3 mm of the cauliflower surface. [Alternate: Whole class works together to remove a total of about 220 g of the outer 2-3 mm of the cauliflower surface. See pictures under the “Results” hyperlink.]

1. Place the cauliflower tissue in a chilled mortar with 40 mL of ice-cold mannitol grinding medium and 5 g of cold purified sand. Grind the tissue with a chilled pestle for 4 minutes (on ice!). [Alternate: Whole class puts the entire 220 g of cauliflower tissue into the kitchen blender. 300 mL of the ice-cold mannitol grinding medium is poured on top. The blender is “pulsed” several times to break up the largest chunks of cauliflower tissue. The cauliflower is then homogenized at “liquefy” speed for 1 minute. See pictures under the “Results” hyperlink.]

1. Filter the suspension through four layers of cheesecloth into a beaker (also wring out the juice); measure and note the volume, then save a small measured amount (~ 2-3 mL) in two microfuge tubes for assays and microscopy. Label the “crude” fraction. Transfer the rest to a chilled 40 mL centrifuge tube. Keep the centrifuge tubes on ice until all groups are ready to centrifuge.

1. Centrifuge the filtrate at 600 x g (4200 rpm, JA20 rotor; 4000 rpm, SS34 rotor) for 10 min at 4° C. Make sure that the centrifuge tubes are balanced; the tubes opposite each other should have the same total volume. Place a 50 mL graduated cylinder on ice. The pellet from this centrifugation is the nuclear pellet.

1. Decant the supernatant from the centrifugation into the chilled graduated cylinder (save the nuclear pellet on ice). Measure the supernatant volume and pour the supernatant into a clean 40 mL chilled centrifuge tube. Spin at 10,000 x g (17,000 rpm, JA20 rotor; 16,000 rpm SS34 rotor) for 20 minutes at 0° - 4° C. Again, be sure that the centrifuge tubes are balanced. Meanwhile, resuspend the nuclear pellet in 5 mL mannitol assay medium. Put another 50-mL graduated cylinder on ice.

1. After the 20 minute centrifugation, transfer the supernatant to the graduated cylinder; note the volume and transfer to a clean tube (Label “soluble” fraction). Resuspend the mitochondrial pellet in 8.0 mL of ice- cold Mannitol Assay Medium. (Use a glass rod or a Pasteur pipette; make sure all the clumps are completely dispersed). Label “Mitochondria”. Store all tubes on ice until the end of the period.

1. You now have 4 samples: crude, nuclear, soluble, mitochondrial.

1. Prepare wet mounts of each and examine by microscopy. Capture a representative image at 400X of each preparation for your notebook. Record observations of the kinds of particles present and their relative frequencies.

1. Prepare slides of all four preparations with methyl green pyronin:

• Place a small drop of each fraction on a clean slide

• Add a drop of stain

• Cover with a coverslip

• Observe at 400X with bright field: nuclei should stain green, cytoplasm red or pink, and mitochondria can be seen as small dots.

• Capture an image of all four preparations for your Results Section

2. Measure the protein concentration in each aliquot as described below and record the concentrations in your notebook. You will need the protein concentrations later to report the specific activity of your preparations.

3. After observing each fraction microscopically and measuring the protein concentration of each fraction, you will need to freeze the fractions for use next week. Put about 1 mL of the crude, nuclear, soluble, and mitochondrial fractions into labeled 1.5 mL microfuge tubes. Put ALL of the remaining mitochondrial suspension (about 7.0 mL) into a labeled sterile 10 mL plastic tube with a screw cap. This larger amount of the mitochondrial fraction will be needed for activity assays next week. Write the names of all group members as well as the day and time of your lab on the outside of a small Ziploc bag. Put all your labeled samples in the small Ziploc bag. Place all of the small Ziploc bags into a large Ziploc bag labeled with the day and time of the lab. Store the large Ziploc bag in the cell biology freezer.

Part B: Determination of Protein Concentration of Your Fractions

Standard Curve for Protein Concentration Determination

Proteins are necessary for cells to function properly. Without them, cells would not be able to have the proper structure, and would not be able to carry out important processes for life. Additionally, the different organelles have different amounts and different types of proteins. Therefore, our different fractions will contain different amounts of protein.

In order to determine how much protein is in each fraction, we must create a standard curve by using protein solutions of known concentrations and determining their absorbance using the spectrophotometer. This standard curve will then be used to determine the protein concentrations in your fractions. A protein that is frequently used for standard curves is Bovine Serum Albumin (BSA). BSA readily dissolves in water to form a colorless solution. Dissolved BSA reacts with specific dye known as Coomassie Blue G-250. Your resulting solutions will turn various shades of blue depending on the concentration of protein that is in your sample. If your sample has a high protein concentration, your solution will turn deep blue. If your solution has a low protein concentration, it will turn a brownish blue. If your solution has no protein in it, then it will be brown.

In this lab you will: A) mix known concentrations of BSA dissolved in buffer with Coomassie Blue G-250, B) measure the absorbance of the resulting blue colored solution and C) plot the measured absorbance of the protein-dye conjugate versus the known concentration of BSA in that sample. The resulting graph will be the protein standard curve that you will then use to determine the amount of protein in your samples of unknown protein concentration.

NOTE: You will be provided with a protein solution containing BSA at a concentration of 100 µg/mL (0.1 mg/mL). This is known protein concentration. You will do serial dilutions of your known protein concentration to make other proteins solutions of known concentration.

1. Make sure that the spectrophotometer has been turned on and is fully warmed up.

1. Label two sets of 13 x 100 glass tubes #1-6a and # 1-6b. (You will do each of the following steps in duplicate).

2. Your BSA working solution is 100 µg/mL. You must now calculate what volume of the BSA working solution you must add to the tubes such that the two sets of six tubes will contain 0, 10, 20, 40, 70, or 100 µg protein respectively. If in doubt, check with the instructor. Write your volumes in the table below.

3. To each tube add the appropriate amount of distilled water to bring the final volume up to 1 mL. All tubes must have the same final volume. Write your volumes in the table below.

4. Add 1 mL of Coomassie Blue G-250 Dye Solution to each tube. Mix by inverting (use parafilm over the top of the test tube), swirling or gently vortexing with a vortex mixer. (Remember, frothing denatures proteins; denatured proteins precipitate, and precipitated proteins are not accurately measured. Only soluble protein is measured).

5. Tube #1 contains no protein so it is considered the "blank”. Pour the contents of tube #1 into a cuvette . Place this cuvette in the cuvette holder labeled “B”.

6. Allow the color to develop for 10 minutes. While the color is developing, zero the instrument using the cuvette in the “B” position.

7. Sequentially place the contents of the other tubes into cuvettes and place them sequentially into the cuvette holders 1, 2, 3, 4 and 5 on the rotating turret. Close the lid.

8. Ten (10) minutes after adding the protein dye solution, measure and record (see the Table below) the absorbance of the cuvettes in positions 1, 2, 3, 4 and 5 at 620 nm. [Note: the color will continue to develop and get darker with time. The standard curve will only be useful when compared to samples of unknown protein concentration that have been similarly exposed to the protein dye solution for 10 minutes.]

9. To create a standard curve, plot the average absorbance at 620 nm (A620) of each pair of samples on the Y-axis versus the µg protein (BSA)/mL on the X-axis. Look at the graph and find the linear region. You will use the linear range to determine the protein concentration in your unknown sample.

|Tube Number |(g BSA to be|Volume BSA |Volume of water |Concentration of BSA |Measured |Average |

| |added |solution to be |to be added |in ug/mL |Absorbance at |Absorbance |

| | |added | | |620 nm |at 620 nm |

|1a |0 | | | | | |

|1b |0 | | | | | |

|2a |10 | | | | | |

|2b |10 | | | | | |

|3a |20 | | | | | |

|3b |20 | | | | | |

|4a |40 | | | | | |

|4b |40 | | | | | |

|5a |70 | | | | | |

|5b |70 | | | | | |

|6a |100 | | | | | |

|6b |100 | | | | | |

Determination of Protein Concentration in your fractions:

Preparation of samples: use one set of each fraction for the protein assay. Dilute your fractions with water indicated in the table below.

1. For a 1/20 dilution, pipette 50 µL of the protein containing cauliflower suspension into a clean cuvette; add 950 µL distilled water and invert to mix. For a 1/100 dilution, add 10 µL of the protein containing cauliflower suspension + 990 µL distilled water. Starting with your lowest dilution for each fraction, then continue with the following instructions:

1. Add 1 mL of Protein Dye Solution to the 1 mL of diluted protein containing cauliflower suspensions. Add 1 mL of Protein Dye Solution to 1 mL of distilled water to serve as your blank.

1. Determine the absorbance of the first dilution for each cauliflower fraction. See a photo of the results of this procedure at:

1. If the absorbance is out of the linear range of your standard curve, then the absorbance reading is invalid. You must repeat the measurement using the greater dilution of your cauliflower fraction.

1. If the absorbance reading is in the linear range of the curve, and above the lowest detectable amount, record this value and use it to determine the protein concentration of that fraction.

1. Using the BSA standard curve, determine the protein concentration of each fraction in µg/mL

2. When finished, rinse all tubes thoroughly in warm running tap water and then submerse them in pan of water for later washing. Cuvettes should be carefully rinsed with warm running tap water and the inside surfaces of the cuvettes should be gently rubbed with wet cotton swabs to remove any adherent Coomassie stain. The cuvettes should be rinsed one final time in RO water and turned upside down to dry on a piece of absorbent material or paper towel.

| |Dilution |Abs 620 nm |[Protein] from standard curve|[Protein] of the original |

| | | | |sample |

|Water (for blank) |NA |0.00 |0 |0 |

|Crude homogenate |1/20 | | | |

|Crude homogenate |1/100 | | | |

|Nuclear fraction |1/20 | | | |

|Nuclear fraction |1/100 | | | |

|Mitochondrial fraction |1/20 | | | |

|Mitochondrial |1/100 | | | |

|fraction | | | | |

|Supernatant |1/20 | | | |

|Fraction | | | | |

|Supernatant fraction |1/100 | | | |

Laboratory #6 Second Week-SDS Polyacrylamide Gel Electrophoresis

Learning Objectives:

1. Learn to do SDS-Polyacrylamide gel electorphoresis, a common technique in Cell Biology

2. Learn how to use stoichiometry to determine how much sample to run on your gel

3. Learn how determine how many proteins are in your individual samples, and how to determine their size in molecular weight

4. Learn how to determine protein content differences between organelles

Experimental Objectives:

1. to separate the proteins in your cell fractions by electrophoresis

2. to stain the proteins for easy visualization

3. to determine the molecular mass of some of the unknown proteins

4. to determine whether the various fractions have unique proteins

Gel electrophoresis is one of the most commonly used techniques in both Molecular and Cellular Biology. In fact, many of you have probably run a gel already. The purpose of doing gel electrophoresis is to separate charged molecule by running them through a matrix (a gel) when subjected to electrical current. The properties of each molecule will determine how far it will run through a gel. Molecules are separated on the gel on the basis of charge, size, and shape, with small molecules migrating further (towards the bottom of the gel) and larger molecules migrating slower (finishing towards the top of the gel).

The most common molecules that Biologists run on gels are nucleic acids (DNA and RNA) and proteins. DNA and RNA are usually run on agarose gels, whereas proteins are most commonly run on SDS-Polyacrylamide gels. However, sometimes DNA and RNA are also run on SDS-Polyacrylamide gels.

Nucleic acids have a negative charge, which helps us in getting them to run on a gel. In order to get the nucleic acids to migrate through the gel, we need to load them orient the electrical field in the correct manner. Each gel has a set of wells at the top, which is where we load our samples. In order to then get them to migrate through the gel, we need to then subject the samples to an electrical field that is properly oriented. To orient the field correctly, we place the negative electrode (cathode) at the top of the gel, and the positive electrode (anode) at the bottom of the gel. When the field is active, the negatively charged nucleic acids will migrate towards the positive electrode, and into the gel, properly separating by size.

Proteins however do not have a negative charge, so how do we get them to migrate through the gel? The simple answer is that we use the detergent SDS (Sodium Dodecyl Sulfate), by placing it both in the gel, the running buffer and the sample buffer. The SDS is a negatively charged molecule and acts as a denaturant, removing both secondary and tertiary structure from the proteins. The SDS molecules that bind the protein also confer a large net negative charge on it, approximately proportional to the molecular weight of the protein. Therefore, when we subject the proteins to our electrical field, the now negatively charged molecules will migrate through the gel matrix toward the anode, with smaller proteins migrating faster, and larger proteins migrating faster. Actually, they will migrate toward the anode at a velocity that is roughly proportional to the log of the molecular weight of each polypeptide. . Since the denatured proteins migrate through the gel on the basis of size, it is possible to estimate molecular weight of individual polypeptides. Standard proteins of known size are electrophoresed on the same gel along with sample proteins

In this lab, we will take the fractions we made from our cell fractionation and run them on an SDS-Polyacrylamide gel. Alongside our samples will run a ladder containing proteins of known molecular weight, which we use to prepare a standard curve. We will then use the standard curve to estimate the molecular weight of various proteins in our fractions.

Materials:

Mini-Protean electrophoresis equipment (Bio-Rad) Coomasie Blue stain

Mini-Protean Ready Gels (12%) (Bio-Rad) Power supply

Cell fractions from previous experiment Micropipettors, tips, microfuge tubes

4x Sample Buffer Boiling water bath

5x Running Buffer (dilute to 1x before use) Unknown protein @ 1 mg/mL

SDS-PAGE prestained standards

Destain I (50% methanol, 5% acetic acid, freshly made)

Destain II (7% acetic acid, 5% methanol, freshly made)

Procedure:

NOTE: The Mini-Protean III is a miniature vertical slab gel unit intended for rapid electrophoresis of protein samples of small volume. Because its gels are small, proteins may be separated in less than 45 minutes. The Mini-Protean III can run two polyacrylamide gel sandwiches simultaneously, if needed. The polyacrylamide gels used in this lab are precast gels (4% acrylamide in the stacking gel and 12% acrylamide in the running gel). The upper, or stacking gel (4% acrylamide) slightly restricts protein migration and serves to concentrate the protein samples. The running gel (12%) serves to separate individual polypeptides into discrete bands. For a general idea of how this looks see supplemental Figure 2.

To visualize the proteins after electrophoresis, the gel must be stained with Coomassie Blue for 30 minutes, and then destained. To handle and to preserve the gel, the gel can be enclosed in a sealable plastic bag with a small amount of Destain II. The gel can be photographed with a digital camera to provide all students with a digital image of their gel.

Each gel will hold up to 10 samples. Each group will use one gel and will arrange their samples asymmetrically so that they can accurately identify lanes after the gel has been stained. Place a drawing of a gel in your notebook identifying the order in which you will be running the samples.

Each Mini-Protein III electrophoresis apparatus will accommodate two gels. Two groups will use one Mini-Protein III apparatus and one power supply. Two groups will electrophorese their gels on one apparatus.

1. Find the small fractions that you froze at the end of the cell fractionation experiment.

2. By now you should have determined the protein concentration of each cell fraction. Now, determine what volume of each sample (crude, nuclear, soluble, mitochondrial) will contain 50 µg of protein. For example, if you calculated that your protein yield was 5 mg protein /mL (5 µg/µL), you would need 10 µL of that fraction to give you a 50 µg concentration. Place the appropriate volume (based on protein concentration) of each sample into a labeled microfuge tube.

3. Add 1/4 volume of 4x Sample Buffer to each sample. For example, if the volume of your protein sample is 10 µL, add 2.5 µL of sample buffer. Also add 2.5 µL sample buffer per 10 µL of your protein standards.

4. Place the microfuge tubes containing your samples and sample buffer in a boiling water bath and boil the samples for 2 minutes. Remove the microfuge tubes and place tubes on ice.

5. Insert the precast gel to the gel apparatus as demonstrated by the instructor.

6. Add 1x Running Buffer to the buffer chambers of the electrophoresis apparatus as demonstrated by the instructor. [Note: About 500 mL Running Buffer must be prepared for each group of students.]

7. It is usually best to avoid the outmost lanes. Load the boiled protein samples into the bottom of a well as demonstrated by your instructor, in the order shown in Table 1.

|Lane 1 |Empty |

|Lane 2 |SDS-PAGE molecular weight standards |

|Lane 3 |mitochondrial fraction |

|Lane 4 |soluble fraction |

|Lane 5 |nuclear fraction |

|Lane 6 |crude fraction |

|Lane 7 |SDS-PAGE molecular weight standards |

|Lane 8 |your choice |

|Lane 9 |your choice |

|Lane 10 |Empty |

1. Connect the leads to a power supply (red to red and black to black), and electrophorese the samples until the bromophenol blue dye front has traveled to the very bottom of the gel (200 V for ~ 45 minutes).

2. After electrophoresis, carefully remove the fragile gel from between the glass plates, and submerge the gel in Coomassie Blue stain. Shake gently on the shaker for at least 30 minutes.

3. Remove the gel from the stain solution and place in Destain I for 15 minutes to 1hr. Remove and put in Destain II 1 - 4 hours until the background is clear. (If you leave it too long in the destain, even the proteins will become destained).

4. Put your destained gel on a piece of saran wrap or in Ziploc bag and photograph it with a digital camera. Include a centimeter ruler in your photograph so that you can easily quantify your measurements. You should put a print of this image into your notebook.

5. Use an image modifying software (eg: Photoshop) to make a grayscale TIF file out of your image and save it as sdsgel.TIF. Save the file somewhere on your computer where you can easily find it when needed.

6. Use Scion Image (NIH Image) to measure the migration distances on of the various proteins on your gel. You will need to analyze the grayscale sdsgel.TIF file. You will need to set the scale in millimeters using the ruler that you included in your photograph. Once the scale is set, start your measurements at the beginning of the resolving gel and end them at the forward edge of the band under consideration. Remember to measure the migration distance of the bromophenol blue for each lane so that you can calculate the Rf for each band. Record all of your data in spreadsheet for easy data manipulation and graphing. You may print a copy of your spreadsheet and put it into your notebook.

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