You will isolate cellular protein from whole tissue using CelLytic MT. This is a proprietary reagent that contains a mixture of salts and detergents to effectively disrupt lipid membranes, lyse cells, and buffer the cellular proteins at the appropriate pH. This solution will also be supplemented with a cocktail of protease inhibitors to minimize the impact of the numerous proteases that are ubiquitous within all cells.
After extraction, you will determine the concentration of proteins in your sample. Due to the high variability of protein structures, these molecules do not uniformly absorb light at any specific wavelength like nucleic acids. We will use the Bradford Assay to determine the concentration of proteins in your sample. This is a colorimetric assay that uses a reagent (Coomassie Blue) that interacts with proteins. When the reagent is mixed with a solution containing proteins, the solution will turn different intensities of blue depending on the amount of proteins present in the sample. This blue dye absorbs at 595nm and the absorbance can be directly correlated to a specific amount of protein present in the sample when compared to a standard curve.
A standard curve is created by conducting the assay with a set of proteins of known concentration (standards). A dilution series is created from these standards to span the maximum possible range of detection for the assay. This dilution series is then processed according to the assay protocol and measurements are taken. The measurements resulting from these standards are then plotted on a graph and a best fit line is created using these data points. From this best fit line, an equation can be obtained for the slope of the line (y = mx+b). This equation allows you to now put in any value you obtain from an experimental sample and determine the concentration of proteins in that sample.
Due to the dynamic nature of the reaction taking place in the Bradford Assay (and virtually all assays), a standard curve should be created using the exact same reagents and equipment that will be used for the experimental samples. This helps to account for variation between different pieces of equipment as well as slight differences between reagent lots. Ideally, a new standard curve should be made each time a set of experimental samples are being assayed to ensure the best accuracy. For the sake of time, a standard curve has already been determined.
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IMPORTANT NOTES
1. Wear clean gloves - Proteases are present on you skin and are detrimental to the integrity of your samples.
2. Mixing - The Bradford assay works best when samples are mixed well. Invert tubes frequently during incubations, and immediately before measuring absorbance to ensure accurate absorbance readings.
PROTEIN EXTRACTION PROTOCOL
Extraction:
1. Add 0.5mL of CelLytic MT solution to a 1.5mL snap cap tube.
2. Using a clean razor blade, cut a piece of frozen tissue weighing 25mg and add to tube containing CellLytic MT solution.*
*to save time, this step has been performed for you.
3. Homogenize the tissue with a disposable pestle.
4. Close the tube and invert the tube several times.
5. Spin the tube in a refrigerated microfuge for 10mins. @ max speed.
6. While spinning, label a fresh tube with the word "Protein", source organism/tissue, your initials, and today's date.
7. Carefully transfer supernatant to labeled tube and store tube on ice.
Quantification:
8. In a fresh 2 mL tube labeled as 'sample', dilute an aliquot of the sample 1:2 by pipetting 15uL of your protein sample and 15uL of DI water and mixing well.
this dilution step is performed to ensure the sample absorbance falls within the range of the standard curve
9. In a second 2 mL tube pipette 30uL of DI water (this tube will serve as your blank). Label tube as 'blank'
10. To both tubes add 1.5mL of Bradford reagent.
11. Invert the tubes several times and then incubate at RT for 10mins.
12. Mix the 'blank' tube and transfer 1mL to a plastic, disposable cuvette.
13. Zero the spectrophotometer using your blank sample
14. Mix the 'sample' tube and transfer 1 mL to a plastic, disposable cuvette
15. Measure the absorbance at 595nm and record the value.
16. Remove the cuvette from the spectrophotometer. Using a P1000 set to 1mL, carefully pipette the solution in the cuvette up and down a couple of times to mix.
17. Measure the absorbance at 595nm and record the value.
18. Average the two absorbance values you recorded.
19. Back-calculate your protein concentration using the standard curve below. Don't forget to account for the dilution in step 8!
Storage:
20. Store your protein sample at -20ºC.
Standard curve was generated per Manufacturers Instructions. Please Read and Understand!
Pierce:Coomassie (Bradford) Protein Assay Kit
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Background - SDS - Polyacrylamide Gel Electorophoresis (SDS-PAGE)
SDS-PAGE is the process of separating proteins from one another on the basis of molecular weight. A mixture of proteins is subjected to an electric field and pulled through a polyacrylamide matrix towards the cathode. However, proteins must be treated prior to separating them in this manner. This is because the charge on any given protein is dependent upon the amino acid sequence as well as the pH of the solution. Thus, a crude protein extract from cells or tissue contains a hetergenous mixture of proteins with varying charges. Without treating them in some manner, the proteins will migrate independent of their molecular weight. Additionally, proteins have various tertiary or quarternary structure that can influence the rate at which they migrate in an electric field. In order to address these issues, protein samples are prepared in a specific fashion to linearize the proteins and impart the same charge to all proteins in the sample to ensure that they become homogenous and their migration rate during SDS-PAGE is solely due to molecular weight.
Protein samples are combined with a reducing sample buffer. This reducing sample buffer contains sodium dodecyl sulfate (SDS), B-mercaptoethanol, glycerol, Bromophenol blue and a buffer. SDS imparts an overall negative charge to all the proteins in a sample. This ensures that all of the proteins will migrate in the same direction (towards the cathode) when placed in an electric field. B-mercaptoethanol is a reducing agent. It accepts electrons from disulfide bonds formed between two cysteine residues. It serves as one step to help break any tertiary or quartenary structure of proteins in the sample. Glycerol simply serves as a sinking agent for your sample. It has a greater density and will allow your sample to sink into the buffer contained in the wells of the gel. Bromophenol blue is a negatively charged dye that allows one to visually track the migration of your samples through the polyacrylamide gel. Bromophenol blue migrates at the same rate of proteins ~5-7kDa. Finally, the buffer is present to maintain the appropriate pH for your sample. Once samples have the appropriate amount of reducing sample buffer, they are boiled. Boiling causes the proteins to fully denature, eliminating any tertiary or quartenary structure and leads to linear chains of amino acids.
Samples are then run through polyacrylamide gels. Traditionally, a single polyacrylamide gel is actually comprised of two gels with different percentages of polyacrylamide, pH and buffer. The top portion of the gel, relative to the bottom portion of the gel, has a lower percentage of acrylamide, a lower pH and a lower concentration of buffer. This gel is referred to as the stacking gel. The bottom gel has higher mounts of all three components listed above and is called the running gel.
The low percentage of polyacrylamide in stacking gels allows all the proteins in the sample, regardless of molecular weight, to quickly and easiy migrate through the gel. When the samples begin to enter the lower gel containing a higher percentage of polyacrylamide (running gel), the proteins are now all "stacked" upon one another. This allows all of the proteins in a sample to enter the running gel at essentially the same time. Additionally, the differences in pH and buffer content between the stacking and running gels leads to a local increase in voltage around the sample, which helps drive the sample from solution in the well into the polyacrylamide matrix of the stacking gel.
After SDS-PAGE is complete (when the dye front has reached the bottom of the gel), the gel is either set up for Western blotting (see Lab #4) or is stained to reveal the proteins. There are a number of stains that can be used, depending on the sensitivity needed to visualize proteins of interest, but we will use Coomassie Brilliant Blue. This is a non-selective stain, meaning it binds all proteins regardless of their amino acid makeup. Additionally, it is cheap and rather sensitive. The dye will initially turn the entire gel blue and even a short exposure (5 minutes) often results in over staining. Thus, it is necessary to destain the gel to wash the dye out of the areas of the gel where no protein is present. After destaining, the proteins should appear as blue/purple bands and the rest of the gel should remain relatively clear.
PROTEIN GEL PROTOCOL - See also Manufacturers Protocol / Manual: Precise™ Protein Gels
1. Begin boiling water on hot plate.
2. Thaw your protein extract from last week. Mix well by inverting tube several times.
3. In a fresh, 1.5mL SCREW CAP tube add 15uL of your protein sample and 15uL of 2X Reducing Sample Buffer.
4. Mix sample by flicking. Briefly centrifuge (10s) to pool liquid in bottom of tube.
5. Boil sample for 5 mins.
6. While sample is boiling, observe assembly of gel box and gels. Rinse gel wells thoroughly as demonstrated.
7. When sample is finished boiling, immediately centrifuge for 1min. to pool liquid.
8. Slowly load your entire sample into the appropriate well using a gel loading tip.
9. Put lid on gel box and plug electrodes into appropriate receptacles on the power supply.
10. Turn power supply on and set voltage to 150V. Run for 45mins.
11. Add ~150mL (does not have to be measured - just need enough to cover the gel) of Coomassie Stain to a designated container.
11. Turn off power supply and disconnect gel box from power supply.
12. Remove lid from gel box.
13. Disengage the tension wedge.
14. Remove gel from gel box.
15. Carefully crack open cassette to expose gel.
16. Trim wells at top of gel.
17. Notch a designated corner of the gel to help you remember the correct orientation of the gel (i.e. which is the top/bottom of the gel, which is the right/left side(s) of the gel)
18. Place gel into container with Coomassie Stain.
19. Incubate on shaker/rocker for 5 mins.
20. Carefully pour stain back into original container. Be careful not to dump out gel!
21. Rinse gel briefly with 10% acetic acid and pour this wash down the drain.
22. Add ~250mL (no need to measure) 10% acetic acid to container with gel. Incubate on shaker/rockers for 15mins. Change out buffer and repeat until bands become clearly visible. This may need to incubate O/N. If so, cover container with plastic wrap and leave on shaker/rocker.
Principles of Western Blotting:
Protein Transfer:
After the protein components have been sufficiently separated by electrophoresis, they can be transferred to a nitrocellulose membrane. The transfer process uses the same principle as SDS-PAGE – this time the electric current is applied at 90 degrees to the gel and the proteins migrate out of the gel onto the membrane. The membrane used for our lab is a positively charged nitrocellulose membrane. Remember from the sample preparation for SDS-PAGE, we've treated the proteins with SDS to impart an overall negative charge to these molecules. During transfer, the proteins migrate through the electric field, out of the gel and onto the membrane. The negative charge of the proteins allows them to efficiently bind to the positively charged membrane. However, nitrocellulose membranse will bind virtually any proteins, regardless of charge. The charge simply ensures a stronger bond. Thus, it is extremely important to handle nitrocellulose membranes with clean gloves and clean utensils to minimize extraneous proteins from being bound to the membrane.
Western Blot Procedure:
• Blocking
The membrane is blocked in order to reduce non-specific protein interactions between the membrane and the antibody. This is achieved by placing the membrane in a solution of non-specific proteins (usually BSA or non-fat milk). The proteins in the blocking solution coat the remaining areas of the membrane where no protein is bound from the transfer. The reason this is necessary is described in the next step.
• Primary Antibody
The first antibody to be applied (specific for protein of interest) is incubated with the membrane. The primary antibody is specific for the protein of interest (in this case HSP70), and, at appropriate concentrations, should not bind any of the other proteins on the membrane. Remember, antibodies are proteins, too. If we had not blocked the membrane, the antibody would end up binding to both the membrane and your target protein. This would result in extremely high background (signals not related to the intended target protein(s)) and would use up a significant amount of the available antibody, making the interpretion of results difficult, if not impossible.
• Secondary Antibody
After rinsing the membrane to remove unbound primary antibody, a secondary antibody is incubated with the membrane. It binds to a species-specific portion of the primary antibody. Due to its targeting properties, the secondary antibody tends to be referred to as "anti-mouse," "anti-goat," etc., depending on the animal species that the primary antibody was created in. This secondary antibody is typically linked to an enzyme that allows for visual identification. In our case, the antibody is linked to an alkaline phosphatase (AP).
• Developing
The unbound secondary antibodies are washed away, and the enzyme substrate is incubated with the membrane so that the positions of membrane-bound secondary antibodies will either change color or emit light. Bands corresponding to the detected protein of interest can be visualized. Band densities in different lanes can be compared providing information on relative abundance of the protein of interest. The kit we will be using for visualization (Invitrogen's WesternBreeze Chromogenic Kit) utilizes an enzymatic reaction that creates a dark purple precipitate that can be seen with the naked eye.
The chromogenic system emplyed in the WesternBreeze Chromogenic Kit is the combination of BCIP (5-Bromo-4-Chlo ro-3'-Indolypho sphate p-Toluidine Salt) and NBT (Nitro-Blue Tetrazolium Chloride). Together they yield an intense, insoluble black-purple precipitate when reacted with Alkaline Phosphatase. The NBT/BCIP reaction is illustrated in the figure below. This reaction proceeds at a steady rate, allowing accurate control of the relative sensitivity and control of the development of the reaction.
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Transfer Proteins to membrane
1. Cool the transfer buffer to 4°C.
2. Soak the filter paper, membrane and gel in Transfer Buffer for 15 minutes.
3. Assemble the blotting sandwich in a semi-dry blotting apparatus as follows:
• Anode (+++)
• Filter paper
• Nitrocellulose Membrane
• Gel
• Filter paper
• Cathode (– – –)
4. Transfer the blot for 30 minutes at 20V.
5. Remove the gel from the sandwich and rinse with transfer buffer.
6. Use a cotton swab to remove any adhering gel from the membrane.
Western Blotting Protocol
Western Breeze Manufacturer's Protocol
General Guidelines
• Avoid touching the working surface of the membrane, even with gloves.
• Work quickly when changing solutions as membranes dry quickly. If the membrane dries, re-wet the membrane with methanol and rinse with water before proceeding.
• Add solutions to the trays slowly, at the membrane edge, to avoid bubbles forming under the membrane. Decant from the same corner of the dish to ensure complete removal of previous solutions.
1. Prepare 20 mL of Blocking Solution
Ultra filtered Water 14 ml
Blocker/Diluent (Part A) 4 ml
Blocker/Diluent (Part B) 2 ml
Total Volume 20 ml
2. Place the membrane in 10 ml of the appropriate Blocking Solution in a covered, plastic dish provided in the kit. Incubate for 30 minutes on a rotary shaker set at 1 revolution/sec.
3. Decant the Blocking Solution.
4. Rinse the membrane with 20 ml of water for 5 minutes, then decant. Repeat once.
5. Prepare 10 mL of Primary Antibody Solution (1:3000 dilution)
Blocking Solution 10 ml
HSP 70 antibody 3.3 µl
Total Volume 10 ml
6. Incubate the membrane with 10 ml of Primary Antibody Solution for OVERNIGHT
NEXT DAY
Decant Primary Ab, saved at 4C.
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Ponceau S stain for Western Blots
This is a rapid and reversible stain for locating protein bands following a western blot.
Here is the recipe:
- 1 g Ponceau
- 50 ml acetic acid (100%)
- Make up to 1 L with water
- 0.1 g Ponceau
- 50 ml 10% acetic acid
- 50 ml dH20
Here is the pdf for the product:
Ponceau-S-Stain.pdf