Unit 3
Unit 4
Analysis of Proteins by Electrophoresis
Introduction:
Electrophoresis is the process of placing proteins in a matrix and applying an electric field. This will separate the proteins according to their size, shape and charge. Gels made from polymerized acrylamide—polyacrylamide gels—are frequently used because they have high resolving power, can use relatively larger amounts of protein, do not interact with the proteins and have a stable matrix. Typically the polyacrylamide is crosslinked with N,N’-methylene-bis acrylamide in a polymerization reaction that is initiated by the molecule N,N,N’,N’-tetramethylethenediamine (TEMED). If low concentrations of acrylamide and bis-acrylamide are used, the pores are larger allowing for analysis of higher molecular weight proteins. Conversely, smaller molecular weight molecules are analyzed in gels with higher concentration of acrylamide and bis-acrylamide. The entire complex forms a matrix of fibers as shown in Fig. 1 below, where represent the polyacrylamide and the represent the crosslinking:
Fig. 1: Matrix of Polyacrylamide with Crosslinking
The PAGE gels are polymerized between two square panes of glass that are sealed around the edges and stood upright during polymerization. The resulting gels are run upright as well, giving rise to the common name of “vertical gel electrophoresis”.
PAGE gels may be continuous or discontinuous. Continuous gels have a gradient of acrylamide and bis-acrylamide concentration with the most concentrated gel at the bottom of the gel.
More often, a discontinuous gel is used. In this gel the lower 80 percent of the gel is called the resolving gel. This gel usually contains a higher percentage of crosslinks and is poured first. On top, there is a stacking gel that contains fewer crosslinks and prepared with a buffer containing fewer mobile ions. A comb is inserted when pouring the stacking gel to produce the wells into which the samples are placed. The stacking gel has lower resistance than the resolving gel so proteins will travel quickly through the stacking gel and concentrate at the interface. This allows the proteins to enter the resolving gel close together and therefore increases the resolution.
Native gels
When an electrical field is applied to the proteins they will travel down the gel. Their movement is influenced by many factors: size, shape, and charge. Smaller proteins will move more quickly through the gel than larger ones due to less frictional drag. For the same reason, tightly coiled globular proteins will travel more quickly than more loosely packed proteins or extended structure fibrous proteins in an electrophoretic gel. The ratio of charged amino acids in a protein will also influence movement through the gel; those proteins with more negatively-charge (acidic) amino acids than positively-charged (basic) amino acids will travel more quickly towards to positive anode due to their large net negative charge. A protein with a net positive charge will, in fact, migrate in the wrong direction from the sample well, away from the anode. Native gels when the conformation of the protein needs to be preserved, usually because the location of the protein of interest must be identified by its structural function. For example, if an enzyme must be identified by its enzymatic activity, a native gel will be used. Also, Western blots are typically performed with native gels to ensure that the antibody detection of a protein of interest will work.
Denaturing gels (SDS-PAGE)
With all these factors influencing the movement of native proteins through the PAGE (polyacrylamide gel electrophoresis) gel, it is nearly impossible to predict where a given protein will migrate or to analyze the results of a native gel. In order for proteins to behave similarly, they must be denatured into a uniformly extended shape. That way the unfolded proteins will behave similarly to nucleic acids in that the short polypeptides travel further down the gel and the longer polypeptides do not travel as far. Unlike DNA molecules, however, a denatured protein does not have a uniformly-charged structure. In order for denatured proteins to move similarly to each other in an electrophoretic gel, they must have a uniform charge-to-mass ratio.
There are two protein denaturing agents that are commonly used together in PAGE electrophoresis: a sulfhydryl reducing agent and an ionic detergent. Ditihiothreitol and 2-mercaptoethanol are short molecules containing a sulfhydryl (-SH) group that will break up the disulfide bridges and help to desrupt any tertiary structure and/or quaternary structure of the proteins due to disulfide bridges. (The sulfhydryl group is what gives these compounds their “rotten egg” smell.)
The main denaturing agent is an ionic detergent, sodium dodecyl sulfate or SDS. This detergent is the sodium salt of a 12-carbon alkyl sulfate compound. SDS actually serves two functions. First it disrupts the secondary, tertiary, and quaternary structures of the protein by breaking the ionic and hydrogen interactions between the amino acids of the protein, as well as interfering with the hydrophobic interactions responsible for correct folding of the protein. Secondly, SDS will coat the denatured proteins’ hydrophobic amino acid side groups with its hydrophobic docedyl tail, thereby coating the protein with negative charges from the sulfate head group of the detergent.
Typically, 1% SDS and 0.1 M mercaptoethanol are used along with high temperatures to completely denature proteins and coat their extended structures with negative charges. This gives rise to the SDS-PAGE electrophoresis that is the most common method of identifying an unknown protein or determining its molecular weight. In order to aid with this, a set of proteins markers of known molecular weight are commonly run along with the unknown. Analysis of sizes of protein bands in the gel is then a straightforward comparison to the migration distances relative to the molecular weight markers.
The samples of protein are diluted with a “sample buffer.” In addition to the the mercaptoethanol and SDS denaturants, the sample buffer contains a tracking dye that will travel with the front and determine how far the gel has run. Glycerol is also included for increased density of the sample in order for it to settle into the bottom of the well and not go floating off into the electrophoresis buffer. Typically somewhere between 10 and 40 (g protein are loaded into a well, depending on the purity of proteins in the sample. This total volume is typically about 20-40 (L, half of which is protein and half is the 2X sample buffer.
Lab 4-A:
Polyacrylamide Gel Electrophoresis
Introduction:
In the beginning of this unit there was a brief discussion of polyacrylamide gel electrophoresis and the different components used. Review this before coming to class.
Tonight we will be running a native gel and a SDS-PAGE gel on the purified green fluorescent protein (GFP) samples from Unit s 2 and 3. We will load two identical gels with identical samples, but samples with be prepared and run under two conditions: denaturing and nondenaturing. Once the gels are run, we will examine both under a UV lamp to look for fluorescence by the GFP band. We will then stain all of the protein bands with Coomassie blue, a stain for proteins. The Coomassie blue will stain the gel as well as the proteins, so the finishing step is destaining to remove the Coomassie blue that did not react with the protein bands.
|Safety: |
|1. The wires connecting the cell to the power supply must be in good condition, not worn or cracked. Broken or worn wires not only cause |
|rapid changes in resistance that adversely affects electrophoresis, but they also create an electrocution hazard. |
|2. Make sure the area around the power supply is dry. |
|3. The area for at least 6 inches around the power supply and cell should be bare of clutter and other equipment. Clear space means any fire|
|or accident can be more easily controlled. |
|Wear gloves while loading and handling the gels; the unpolymerized acrylamide is a neurotoxin. |
|5. Coomassie blue will stain clothing and hands. In addition it is very acidic. Wear gloves when handling the staining and destaining |
|solutions. Lab coats or aprons are recommended. |
| |
Protocol:
Part I: Preparing solutions
Students may collaborate in the preparation of the following solutions for use by the entire class. Part of this collaboration is the sharing of the Media Prep Form describing the preparation of each solution in for individual lab reports. Each prepared solution should be labeled with its contents and concentrations, as well as a control number that cross references the solution to its Media Prep Form. Students using a solution should record its control number in their lab report.
Part II: Preparing the gels
1. With gloved hands, carefully un-wrap the cellophane from two PAGE gels, which are enclosed in a gel support chamber. The gel support chamber consists of thin plastic plates sandwiching the gel. One gel will be run under non-denaturing conditions, while the other will be run under denaturing conditions.
2. Handle the support chamber gently to avoid distorting the gel within. Drain any excess water out of the wells (the indentations in the top of the gel), and then clamp the gel support chamber onto the electrophoresis apparatus according to the instructions provided.
3. Pour the appropriate running buffer (see table below) into the top half of the apparatus to check for leaks. If sufficiently sealed, no buffer will appear in the bottom chamber. If this is the case, continue to fill the top of the apparatus until the gel and electrodes are covered with buffer.
|Component |Concentration in the SDS-PAGE running buffer|Concentration in the native gel PAGE running |
| |(1X) |buffer (1X) |
|Trizma base |25 mM |25 mM |
|Glycine |192 mM |192 mM |
|SDS |0.1% |----- |
NOTE: The final pH of the running buffers should be 8.3.
4. Gently remove the comb from the stacking gel.
5. Rinse any unpolymerized acrylamide from the wells by pipetting the running buffer in and out of each well. The wells are too thin to use micropiptter tips; you will need to use special gel loading tips. Take care not to puncture the sides or the bottom of the well with your pipetter tip. This takes a steady hand – it may help to support the micropipettor with your other hand and to support your elbows on the lab bench top.
Part III: Preparing and loading samples into the gel wells
1. Defrost your samples of GFP from Labs 2 and 3.
2. Label microfuge tubes and add 20 (L of each GFP sample with an equal volume of 2X sample buffer. Choose GFP samples to represent different stages of your purification process. If purified GFP eluted from your column in multiple fractions, select the fractions that had the greatest amounts of GFP fluorescence. The components of the 2X sample buffer are given below. You need 11 x 20(L of each for two 10 well gels.
|Component |Concentration in the 2X SDS-PAGE sample |Concentration in the 2X nateive gel PAGE sample |
| |buffer |buffer |
|Tris-Cl pH 6.8 |126 mM |126 mM |
|Glycerol |20% |20% |
|Bromophenol blue |0.0025% |0.0025% |
|SDS |4% |----- |
3. Pipet up and down to ensure mixing. If your protein MW standards are not in sample buffer, add 10 uL to an equal volume of 2X sample buffer as well.
4. Incubate SDS-PAGE samples and protein MW standards in a 60° C water bath for 5 minutes. Dry the tubes with a paper towel and place them in the microcentrifuge in balanced positions. Centrifuge for 5 sec to force the solutions to the bottoms (simply turning the centrifuge on and immediately turning it off is sufficient.)
5. The wells are identified as #1 through #10, starting with #1 on your left as you face the front of the gel.
6. Read the Good Laboratory Practice Tips (below) and load 30 (L of each sample into a separate well and record the contents of each loaded well. Be sure to use a new gel loading tip for each sample loaded. Load each of the two gels in the same order.
| |
|GLP tips: |
| |
|To load a sample into a well, |
|Adjust an automatic pipetter to deliver the correct amount of the sample |
|Attach an ultrathin gel loading tip. |
|Withdraw the correct amount of your sample from your microcentrifuge tube and insert the pipetter tip into the top of the well to at least |
|four mm of the bottom of the well. Take care not to puncture the sides or the bottom of the well with your pipetter tip. This takes a steady|
|hand – it may help to support the micropipettor with your other hand and to support your elbows on the lab bench top. Make sure that the |
|pipette tip is between the glass sandwich of the gel and very slowly and gently expel the solution from the pipetter tip into the well while |
|holding the pipetter steady. The blue solution should fall to the bottom of the well, gradually filling it. |
|Do not press the pipetter to the second stop – it is important to avoid blowing air bubbles into the well. |
|Do not release your thumb until you have slowly withdrawn the pipetter tip from the well, so that you avoid removing the sample that you have |
|so carefully loaded! |
| |
|Troubleshooting: |
|If the sample overflows into the adjacent well, you may be trying to load too much sample. Alternatively, you may be expelling the sample |
|with too much force, or not withdrawing the micropipettor tip enough to make room for your sample as it fills the well. You may quickly |
|withdraw any sample that has overflowed into an empty well. |
|Work quickly to minimize the diffusion of samples from the wells or between wells. |
| |
7. Place a new tip on the automatic pipetter, then follow the procedure just described to load your “MW standards” into one well of each gel.
Part IV. Running your gels
Check to make sure that the upper buffer is not leaking into the lower chamber. If it is, you must disassemble the apparatus and fix the leak with vacuum grease along the rubber sealing ring. If the upper buffer chamber is not leaking, add running buffer to the lower chamber until the bottom of the gel is submerged.
□ Electric shock hazard! Be sure to follow instructions exactly!
1. Wipe up any spills with paper towels so that your work area is dry.
2. Connect the terminals, turn on the power supply and set the voltage to 150 V. Run the gel for 45 – 60 minutes until the dye front is near the end of the gel.
3. Plug in the power supply, turn it on, and adjust the voltage to 150V. (It may be hard to get it to stay at exactly 150V, but get as close to that setting as possible.)
4. As demonstrated by your instructor, connect the electrophoresis apparatus to the power supply and plug it in. Have your instructor check your set-up and connections. When given the OK by your instructor, turn on the power supply. Record the time when the power was turned on.
5. Running time is generally between 50 minutes and one hour. As current flows through the gel, proteins that are negatively charged will be pulled towards the bottom of the gel. You will not be able to see the proteins since they are colorless, but the blue tracking dye should remain visible. A few minutes after the power is applied, the blue bands should concentrate as a thin line below each well at-the interface between the stacking gel and the separation gel. The blue bands should then move slowly down through the separation gel gradually becoming thicker as they move downward.
When the blue dye is within 2-3 millimeters of the bottom of the gel, turn off the power supply and unplug it.
6. Wearing gloves, remove the cover from the electrophoresis apparatus. With gloved hands, carry the apparatus to the sink, secure the central section with your thumbs, and invert the apparatus to discard the buffer from the two reservoirs. Carefully remove the gel sandwich from the apparatus. Place on a glass tray or a piece of plastic wrap.
Part V: Staining and Destaining your gels
1. Place a piece of plastic wrap on your table. Remove the clamps holding the “gel sandwich” in place, and lay the gel sandwich on the plastic wrap.
2. Examine your gels under a UV lamp to locate the GFP bands (if any) in each of your gel lanes. Record your observations.
3. To extract your gel from its cassette, find a place to insert a spatula between the two plates where the gel will not be damaged. The stacking gel at the top will be removed from the gel and damage there will be unimportant. Pry the plates apart to open the cassette.
4. You should see a faint seam between the stacking gel (on top) and the resolving gel (on the bottom). Cut along this seam gently with a scalpel blade and scrap away the stacking gel while leaving the resolving gel behind.
5. With one of the plastic plates removed, check for GFP bands again under the UV lamp and record these results.
6. The gel must be delivered to a Coomassie Blue staining solution in order to stain the protein bands.
The gel is very thin and fragile, and you must take special care not to tear it. Use a dH20 squeeze-
bottle to wet the gel. Tilt the plate with the gel at an angle upside down over a large Petri dish
containing Coomassie Blue dyeing solution, placing the gel immediately above the solution. Using a
spatula, gently peel the gel from the plate starting with the corner furthest away from the dye solutions
in the Petri dish. Gently pull the corner of the gel away from the plate until it begins to peel away on its
own due to gravity. Once the gel starts peeling from the plate it should continue to tear itself from the
plate and into the solution without any help from you. Make sure that the gel is lying flat in the staining
solution, adding more staining solution if necessary in order to remove any folds. Cover the Petri dish,
and use a piece of tape to label it with your group name.
7. Place the dish on a rotary agitator and allow it to agitate very slowly for 20-30 min.
8. While your gels are staining, rinse out the electrophoresis apparatus with tap water and then with dH2O, and turn upside down on a paper towel at your work station to drain.
9. The Coomassie Blue stain contains 50% methanol and 10% acetic acid in addition to the dye and can be reused several times, so when you are finished staining, pour the stain solution through the filter-lined funnel into the discard bottle. Carefully hold your gel in the Petri dish with your gloved hand so that you do not lose. Pour out as much of the stain from the Petri dish as possible.
10. Deliver several milliliters of dH20 from a squeeze bottle to the Petri dish containing the gel, swirl briefly to rinse, and again holding the gel in the dish with your gloved hand, discard the rinse water into the sink.
11. Pour "Fresh Destaining Solution" into the Petri dish containing the gel until it is half full. This solution is 50% Methanol + 40% dH20 + 10% Acetic acid. Return the Petri dish to the agitator and record the time when destaining was begun.
12. After 15 minutes of destaining, bring the Petri dish to the sink and pour the liquid into the bottle labeled "Used Destaining Solution”. Pour "Fresh Destaining Solution" into the Petri plate.
13. After 15 minutes, pour the destain from the Petri dish into the bottle labeled "Used Destaining Solution." Ask your instructor to examine the gel and determine if it requires additional destaining. The proteins within the gel should appear as blue horizontal bands in otherwise colorless lanes.
If it is sufficiently destained, fill the Petri dish containing the gel about half-full with dH2O
14. Tape the Petri dish closed, and make sure it is labeled to identify your group. Give the dish to your instructor, who will store it until your next lab period.
Bio-Safe Coomassie Stain-preferred method
1. Rinse gels well (a couple of times) with dH20 to wash off residual running buffer. Any residual SDS may cause the background to stain.
2. Pour bio-Safe Coomassie Stain over gels just until the gel is barely covered. Be sure the gel is in a microwave-safe container, then microwave for 30 seconds. This allows the stain to work faster. Do not overheat the gels!
3. Place the dish on a rotary agitator and allow it to agitate slowly for at least 20 min. You may also shake the gel dish by hand. You may also let the gels stain overnight, then destain for a few hours the next day.
4. Pour out the stain into the sink and flush with plenty of water. Then rinse the gel well (a couple of times) with dH20, and let sit in dH20 for 20 min or overnight. Cover with plastic wrap if leaving overnight. Once the bands are clearly visible and the background is in good contrast, you may take a picture of the gels.
Clean up
Begin to clean up while gels are running, staining, and destaining.
Be careful not to lose any of the gel apparatus, including the plates, the template, the spacers, and the comb. Wash them in soapy water, rinse with tap water and dH2O and leave on absorbent toweling to dry.
Dispose of your excess solutions as instructed by instructor.
Remove label tape and any marks made with a marking pen from all glassware. Wash and rinse all used glassware, give it a final rinse with dH2O, and leave it inverted at your work area in order to drain.
All disposable glassware goes into the special glass disposal receptacle.
Lab 4-B
Analysis of the SDS-PAGE Gel
Introduction:
The GFP protein usually shows up as a large band at 54,000 daltons. Because this is a dimer, there is often a band at 27,000 daltons of the monomers. To determine the MW of an unknown protein band on a gel, its migration distance is compared with the migration distances of the MW standards in the same gel. The gel must essentially be standardized before the size of unknown proteins can be estimated. In this lab we will be using Excel to compile the migration distance values of the gels and draw a graph to be used to locate the band or bands of GFP.
Excel is extremely useful in the lab for handling data. Working in Excel requires mastering some basic concepts and terms. But once mastered, Excel may be used in many applications with only small variations. Tonight we will be learning basic Excel. First you will use data that is given to you. Then you will take a photo of a gel to practice using Excel starting from scratch.
There is one very important rule in Excel: the computer does not think; you are the only one with a brain. You are the one who is manipulating the data and the computer is following your orders. So if you enter a point wrong or give the wrong instruction, the data will be incorrect. Just because a value has a label attached to it and is presented on a spreadsheet does not mean that that value has any meaning. The data and analysis is only as good as the person at the keyboard. So always “eyeball” data and hold it up to scrutiny. Does this piece of data fit in to the other data; does this data make sense? This is very important when working in a lab. Even the best raw data in the world becomes garbage if it is not handled properly.
With that in mind, here is a very brief glossary of some basic terms used in Excel:
□ Sheet: a page in Excel. The standard format opens a new file with 3 sheets. New sheets may be added as needed. Usually each experiment will have its own sheet.
□ Cell: A basic unit of a table or worksheet. It may contain text, a number or a formula. Each cell is named by its position in the column, marked A, B, C, etc. and row, marked 1,2,3, etc. For example, cell A1 is the first cell in column A, row 1.
□ Absolute cell: A cell address in a formula that does NOT change when copied to another cell. An absolute reference has the form $A$1.
□ Relative cell: In formulas, a reference to the address of another cell in relation to a cell that contains a formula.
□ Formula: a mathematical equation such as y = mx + b.
Safety: Wear gloves when handling the gels.
No special safety considerations are necessary when on the computer.
Protocol:
Part I: Practice Exercise:
Below are the molecular weights for the standards on a protein gel. (Make sure this matches the standards that you actually used in the lab!) Included is the migration distance each of the standard bands traveled in mm on that gel. Lastly, there is an unknown band.
| | |Migration distance|
|Protein |M.W. |(mm) |
| |(daltons) | |
| | | |
|Marker 1 |94,000 |9 |
|Marker 2 |66,000 |19 |
|Marker 3 |55,000 |24 |
|Marker 4 |41,000 |34 |
|Marker 5 |36,000 |38 |
|Marker 6 |25,000 |50 |
|Marker 7 |21,000 |58 |
| | | |
|Unknown | |43 |
1. Go to Excel and create 4 columns labeled "Protein," “M.W.,” “log M.W.,” and “mm”. Enter the data above into the appropriate columns on your Excel sheet.
2. Calculate the log M.W. for the markers. Do not put down the unknown yet. When graphing electrophresis data, we must use the log MW on the y-axis in order to obtain a straight line that can be used to calculate the unknown molecular weights. Therefore you will typically see electrophoresis data presented as log MW. Occasionally the data will be plotted on semi-log paper which accomplishes the same thing. To calculate the log MW, highlight the first cell in the log MW column. Type the command: =LOG(mw cell). The mw cell is the comparable cell in the MW column. Then ENTER.
For example, highlight cell B2 and type =LOG(A2), then ENTER to get: 4.97313. Note that you could highlight B2 and type =LOG(94,000) to get the same answer. However, you are almost always better off, selecting a cell rather than the value in that cell. Then if you change the value in the cell, the log will change as well.
Column: A B C
| | | |Migration distance |
|Row: 1 |(MW) |(Log MW) |(mm) |
|2 |94,000 |4.97313 | 9 |
|3 |66,000 | |19 |
|4 |55,000 | |24 |
Etc
3. Repeat for each of the molecular weight values. You may copy the B2 cell and copy it into the other cells of the B column to quickly repeat the calculation throughout.
4. Now you are ready to plot.
Click on the plot icon (the blue, yellow and red bar graph).
Choose xy scatter plot
Choose the upper left curvy subplot
Click on NEXT
Click on Series
In Name, type “Practice Gel”
Highlight x values, then go to your data and highlight the migration distance column
Highlight y values, then go to your data and highlight the log MW column
Highlight Series 2 and press remove.
Highlight Series 3 and press remove.
Click on NEXT
“Practice Gel” should appear in name.
Type “migration distance (mm)” under value x to label the x axis
Type “log MW” under value y to label the y axis
Click on FINISH
Click on AS OBJECT IN to put the graph on your data sheet.
At this point you should have a very nice graph of a straight line going from upper left to lower right. Both axes should be labeled and the title will be on the top. The graph will be in the middle of your data sheet.
Eyeball the graph to make sure that a given point for log MW is really the Rf you calculated for that marker.
4. Now to calculate the MW for the unknown. In order to get this value we must first calculate the slope and the y-intercept for our data. For slope:
Type “slope =” in a lower cell and highlight the cell next to it
Click on the fx icon
Click on statistical and scroll down the function name column to SLOPE
Click on SELECT
Move the box out of the way of your data
Click in KNOWN_Y’s and highlight the log MW column in your data
Click in KNOWN_X’s and highlight the migration distance column in your data
Click OK
The highlighted box next to slope = should give a negative number.
5. For y-intercept
Type “y-intercept =” under your “slope =” cell
Click on the fx icon.
Click on statistical and scroll down the function name column to INTERCEPT. Then OK.
Move the box out of the way of your data.
Under y values, highlight your log MW column
Under x values, highlight your migration distance column
Click OK
6. To get the unknown MW:
Type “Unknown”
Underneath type “mm =” and in the next cell type the value from the table.
Underneath, type “log MW” and in the next cell type
=slope cell * migration distance cell + y-intercept cell
Type “Unk MW =” and in the next cell type =power(10,logMW cell) then
ENTER
This will give the unknown MW. Check whether this MW is correct by estimating from the table.
8. Alternative method:
Right click on the graph
Click on Trend Line
Click on Options
Click on Display to put equation on the graph including the slope and y intercept.
9. Print out your graph and your spreadsheet to hand in.
Part II: Obtaining data from the GFP gels
Follow the newest SOP for the gel doc system.
1. Place a piece of plastic wrap on top of a white light box and gently transfer your gel to the plastic wrap to visualize the gel. If you don’t have a light box, transfer your gel to plastic wrap on a sheet of plain white paper.
□ Transparency film used for overhead projectors cut to a size slightly larger than your SDS-PAGE gel works well for fishing these gels out of a Petri dish without tearing them.
□ You can also use two pieces of transparency films to sandwich your gel for long term storage by sealing the “sandwich” with tape to keep the gel from drying out. Make sure there is a slight excess of destain solution before sealing the edges of the sandwich with tape. This can be photocopied and stored under refrigeration in a ziplock plastic bag for years.
2. Place the gel so the cut corner is to the lower right, which orients the gel in the same way it was oriented when you cut off the corner. Recall that the proteins migrated down through the gel, so the top edge represents the origin (start) of separation.
3. Photograph the gel placed on top of a light box so that the migration distances of each stained protein band may be recorded. Alternatively, the gel in a transparency film “sandwich” can be photocopied with a “photo” setting on the photocopier.
4. During SDS-PAGE, the smallest proteins generally migrate through the gel fastest. Therefore, the proteins which are farthest from the origin have the lowest molecular weight. Measure the distance from the origin (the top of the resolving gel) to the middle of each protein band in the lane containing the molecular weight standards. Make a table listing the molecular weights of each of the protein standards. Based on the known molecular weights of the proteins in this lane, record the migration distance of each MW Standard in a column of your table.
□ You may be able to see the bands more clearly if you place your gel on a light box.
5. Repeat the exercise in Part I for the data to produce a spreadsheet and graph of the MW standards in you gel. Calculate the expected migration distance for GFP (MW = 27,000).
6. Locate the GFP at the expected migration distance in your gel. Look also for a GFP dimer in your gel. Record your observations of the GFP band(s) and any other protein bands in each of the lanes in your gel.
7. Print out your graph and your spreadsheet for your report.
Questions for Unit 4
Lab 4-A:
Read Chapter 27 “Introduction to Bioseparations” by Seidman & Moore in Basic Laboratory Mehods for Biotechnology, section III. “Choosing Bioseparation Methods”, Part C. “High-Resolution Purification Methods” on pp 582-584. Answer the following questions.
a. What are some reasons that PAGE is a technique more appropriate for analytical, rather than preparative, work?
b. Why are proteins preferentially separated on polyacrylamide gels, rather than agarose gels?
c. What is the approximate ratio of SDS molecules per amino acid residue of a protein during separation by SDS-PAGE?
1. Explain the difference between a stacking gel and a resolving gel. Which one has fewer cross links? Why does this gel have fewer cross links?
Lab 4-B:
1. How long did your gel electrophorese? How can you speed up the electrophoresis? What limits the speed that a gel can be run?
2. Give two safety considerations when performing electrophoresis and explain why.
3. Comment on your electrophoresis results.
a. Did you get results that you expected?
b. Did you observe monomers of GFP? How did you know?
c. How many other protein bands did you observe in each lane?
d. In which lanes was the GFP band the predominant protein band?
e. How well did chromatography purify GFP?
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