20.109(S16):Begin Western blot protein analysis and choose system conditions (Day2)

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20.109(S16): Laboratory Fundamentals of Biological Engineering

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Introduction

Previously you learned more about the two cell lines that we will be using during Module 2, and seeded a known quantity of each cell type in preparation for protein analysis. The protein analysis that you will complete serves two important research purposes: 1. The DNA-PK mutant is a negative control and 2. When compared to the wild type, the mutant may give insight into the role of DNA-PK in NHEJ dsb repair.

Today you will lyse the cells you seeded, isolate the protein fraction from the cell lysate, and separate the proteins on a polyacrylamide gel. You will also begin the Western blot procedure by transferring the separated proteins from your polyacrylamide gel onto a membrane. This step will enable you to 'probe' the protein fractions isolated from M059K and M059J for your protein of interest, DNA-PK. To probe the membrane, the teaching faculty will incubate it with an antibody specific to DNA-PK. This antibody is the primary antibody because it binds directly to the protein of interest. During the next laboratory session you will add a secondary antibody. The secondary antibody binds to the primary antibody and provides a means for visualization - in our experiment, the secondary antibody produces a fluorescent signal.

The ability to bind specific proteins using antibodies, or immunoglobulins, is critical in Western blot analysis. Antibodies are typically 'raised' in mammalian hosts. Most commonly mice, rabbits, and goats are used, but antibodies can also be raised in sheep, chickens, rats, and even humans. The protein used to raise an antibody is called the antigen and the portion of the antigen that is recognized by an antibody is called the epitope. Some antibodies are monoclonal, or more appropriately “monospecific,” and recognize one epitope, while other antibodies, called polyclonal antibodies, are in fact antibody pools that recognize multiple epitopes. Antibodies can be raised not only to detect specific amino acid sequences, but also post-translational modifications and/or secondary structure. Therefore, antibodies can be used to distinguish between modified (for example, phosphorylated or glycoslyated proteins) and unmodified protein.

Monoclonal antibodies overcome many limitations of polyclonal pools in that they are specific to a particular epitope and can be produced in unlimited quantities. However, more time is required to establish these antibody-producing cells, called hybridomas, and it is a more expensive endeavor. In this process, normal antibody-producing B cells are fused with immortalized B cells, derived from myelomas, by chemical treatment with a limited efficiency. To select only heterogeneously fused cells, the cultures are maintained in medium in which myeloma cells alone cannot survive (often HAT medium). Normal B cells will naturally die over time with no intervention, so ultimately only the fused cells, called hybridomas, remain. A fused cell with two nuclei can be resolved into a stable cell line after mitosis.

Generating monoclonal antibodies.


To raise polyclonal antibodies, the antigen of interest is first purified and then injected into an animal. To elicit and enhance the animal’s immunogenic response, the antigen is often injected multiple times over several weeks in the presence of an immune-boosting compound called adjuvant. After some time, usually 4 to 8 weeks, samples of the animal’s blood are collected and the cellular fraction is removed by centrifugation. What is left, called the serum, can then be tested in the lab for the presence of specific antibodies. Even the very best antisera have no more than 10% of their antibodies directed against a particular antigen. The quality of any antiserum is judged by the purity (that it has few other antibodies), the specificity (that it recognizes the antigen and not other spurious proteins) and the concentration (sometimes called titer). Animals with strong responses to an antigen can be boosted with the antigen and then bled many times, so large volumes of antisera can be produced. However animals have limited life-spans and even the largest volumes of antiserum will eventually run out, requiring a new animal. The purity, specificity and titer of the new antiserum will likely differ from those of the first batch. High titer antisera against bacterial and viral proteins can be particularly precious since these antibodies are difficult to raise; most animals have seen these immunogens before and therefore don’t mount a major immune response when immunized. Antibodies against toxic proteins are also challenging to produce if they make the animals sick.

Generating polyclonal antibodies.


For Western blot analysis, a high quality antibody can have a relatively low affinity for its target protein. This is because the target is localized and concentrated on a blot, allowing the antibody to bind using both antibody “arms” thereby strengthening the association. Even an antibody that is loosely bound to the blot under these circumstances may dissociate then re-associate quickly since the local concentration of the target protein is high. The lower limit for protein detection is approximately 1 ng/lane, a value that varies with the size of the protein to be detected and the Western blotting apparatus that is used. For most polyacrylamide gels, the protein capacity for each lane is 100 to 200 μg (that would be 20 μL of a 5-10 μg/μL protein preparation). Thus, 1 ng represents a protein that is approximately 0.0005-0.001% of the total cellular protein (1 ng out of 100,000-200,000 ng). Proteins that make up a more significant fraction of the total protein population will be easier to detect.

In addition to the Western blot analysis preparations, you will learn more about the plasmid reporter construct you will use to measure NHEJ. As you may recall, you will use a plasmid that was engineered by the teaching faculty (thank you Leslie and Maxine!). To ensure that you are familiar with the construct and the assay details, you will think through the design considerations that went into building the NHEJ reporter. Briefly, our reporter assay works as follows: a green-fluorescent-protein-expressing plasmid is cut by a restriction enzyme(s) to produce damaged DNA, then transfected into wild-type and DNA-PK mutant cells, and repaired at some frequency that we evaluate by measuring the green fluorescence of the cell populations.

Protocols

Part 1: Prepare cell lysates

In this exercise you will prepare your samples for Western blot analysis. It is very important that you keep the cell lysate cold throughout the procedure.

  1. You have an ice bucket at your bench with the following pre-chilled items inside: two empty eppendorfs, RIPA buffer, protease inhibitors, and PBS.
    • Label the eppendorf tubes as M059K and M059J (plus your section).
  2. Retrieve your 60 mm dish from the incubator in TC, and hold it at a 30-45° angle in your bucket.
    • You may find it helpful to push the ice such that you build a ramp in the bucket that you can use to hold the plate steady.
  3. Add 2.5 μL of protease inhibitors to your 250 μL of aliquotted RIPA buffer and return this mixture to the ice.
  4. Aspirate the media from each well by holding the tip of the pasteur pipet (be sure to cap the pasteur pipet with a clean yellow tip!) at the bottom of the well.
  5. Add 2 mL of ice-cold PBS to each dish.
  6. Obtain two scrapers from the instructors' bench.
  7. Aspirate the ice-cold PBS – be sure to remove ALL of the PBS after this wash.
  8. Add 100 μL of RIPA lysis buffer to the top of each dish and allow it to run down the dish to the bottom.
  9. Dislodge the cells from the dish by scraping each well with a fresh cell scraper.
    • First tilt the plate back and forth to coat the cells with lysis buffer.
    • Then go from top to bottom 'windshield wiper style' to pool the cells at the bottom of each well in the tilted plate.
  10. Transfer the cells from each dish to the appropriate eppendorf tube.
    • At this step it is important to make sure there are no aggregates or clumps of cell lysate remaining in the dish. Do this by tilting the plate so that light reflects off the bottom. If you see any 'chunks', pipette your lysate back into the dish to dissolve the chunk, then re-transfer everything into the eppendorf tube.
  11. Incubate the eppendorf tubes with your cell lysates on ice for 10 min.
    • Meanwhile, label two fresh eppendorf tubes and add them to your ice bucket to chill for a later step.
  12. Spin the cell lysate at maximum speed in the cold room centrifuge for 10 min to pellet insoluble material.
    • The teaching faculty will show take you to the centrifuge when you are ready.
    • This step is typically referred to as "clearing" the lysate. The pellet at the bottom contains the DNA from the cell, and genomic DNA can get very soupy making it difficult to load your lysate on the SDS-PAGE gel.
  13. Transfer the supernatant to the fresh eppendorf tubes – be careful not to disturb the pellet at the bottom!
    • Keep your samples on ice when not in use.

Part 2: Measure protein concentration

You will now measure the total protein concentration in each cell lysate to determine the volume that you will use for the polyacrylamide gel separation.

  1. Immediately before it is your turn to use the spectrophotometer, add 10 μL of each cell lysate to a plastic cuvette.
    • Prepare a 'blank' by adding 10μL of your leftover RIPA buffer to a cuvette.
    • Be careful not to allow 10 μL of lysate to sit in the cuvettes for more than a couple minutes before starting the next step -- the liquid will evaporate!
  2. Carefully take your cuvettes to the spectrophotometer and add 990 μL of Precision Red reagent.
    • Mix by pipetting up and down a 2-3x without introducing bubbles.
  3. Wait 1 minute, then measure each sample at 600 nm.
    • Use the RIPA sample to blank the spectrophotometer.
  4. Calculate the two stock protein concentrations of your M059K and M059J cell lysate using the following information:
    • 1 absorbance unit = 100 ug protein/mL reagent / cm
    • The path length of the cuvette is precisely 1 cm.
    • Remember to account for the dilution factor!
  5. Next, calculate the volumes of lysate and water required to add 20 - 40 μg of total protein to the polyacrylamide gel in a total volume of 20 μL, per lysate sample.
    • If your concentration is greater than 1μg/μL, use water to make up the remaining volume.
    • If your concentration is less than 1μg/μL for at least one sample, scale both samples down to a lower protein amount, such as 10 μg.
  6. Do not throw away the remainder of your cell lysate! We will store these samples at -80 °C in case your Western blot needs to be repeated.

Part 3: Separate proteins using polyacrylamide gel electrophoresis (PAGE)

You will now prepare your cell lysate samples for separation using PAGE. Two teams will share one gel.

  1. Add 4 μL of 6X Laemmli sample buffer to both of your cell lysates.
  2. Briefly vortex each tube, then centrifuge to collect the samples at the bottom of the tubes.
  3. Put lid locks on the eppendorf tubes and boil for 5 minutes in the water bath that is in the fume hood.
  4. Load your samples according to the scheme below.
  5. In your notebook, document the start and stop time of electrophoresis.
    • The teaching faculty will begin electrophoresis after both groups load their samples.
    • The proteins will be separated using 200 V for 35 minutes.
  6. During electrophoresis, work through Part 5 with your laboratory partner.
Lane Sample Volume to load
1 EMPTY N/A
2 Team 1, M059K 20 μL
3 Team 1, M059J 20 μL
4 HiMark ladder 10 μL
5 EMPTY N/A
6 HiMark ladder 10 μL
7 Team 2, M059K 20 μL
8 Team 2, M059J 20 μL
9 HiMark ladder 10 μL
10 EMPTY N/A

Part 4: Transfer proteins onto nitrocellulose membrane

After the electrophoresis procedure, the teaching faculty will assist you in assembling the transfer cassette according the below protocol. If we run short on time, the teaching faculty will complete this part for you.

  1. Wearing gloves, carefully disassemble the electrophoresis chamber.
  2. Assemble the transfer cassette in the following order:
    • Place the black side of the transfer cassette in a tupperware container with transfer buffer. The transfer cassette is color-coded so the black side should face the cathode (black electrode) and the clear side should face the anode (red electrode).
    • Place a ScotchBrite pad pre-soaked in transfer buffer on the black side of the cassette.
    • Place 1 piece of filter paper on top of the ScotchBrite pad.
    • Place your gel on top of the filter paper.
      • Lightly press out any air bubbles that form between the filter paper and your gel.
    • Place a piece of nitrocellulose membrane on top of the gel.
      • The nitrocellulose membrane is white and should be kept between the blue protective paper sheets until use. Wear gloves when handling the membrane to avoid transferring proteins from your fingers to the membrane.
      • Again, lightly press out any air bubbles that form between the gel and the nitrocellulose membrane.
    • Place another piece of filter paper on top of the nitrocellulose.
    • Place a second ScotchBrite pad pre-soaked in transfer buffer on top of the filter paper.
    • Close the cassette, then push the clasp down and slide it along the top to hold it together.
    • Place the transfer cassette into the blotting tank so that the clear side faces the red electrode and the black side faces the black electrode.
  3. Two blots can be run in each tank. When both are in place, insert a magnetic stir bar and an ice compartment into the tank and fill the tank with transfer buffer.
  4. Connect the power supply and transfer at 100 V for 60 min.
  5. After the transfer is complete, turn off the current, disconnect the blotting tank from the power supply, and remove the transfer cassettes.
  6. Disassemble the transfer cassette to retrieve the nitrocellulose membrane and confirm that the pre-stained standard markers transferred from the gel to the membrane.
  7. Cut the membrane such that each team has a membrane with only their samples, and then transfer each membrane to a separate plastic tupperware.
  8. Add enough Odyssey blocking buffer to the plastic dish to just cover the membrane.
  9. Your membranes will be stored at 4°C.

Part 5: Engineer pMax-EGFP-MCS

In this module you will use an NHEJ reporter to measure dsb repair. The plasmid construct was recently engineered by your teaching faculty and is a new iteration of a blue fluorescent protein (BFP) reporter originally developed in the Samson Laboratory. In this exercise you will familiarize yourself with the NHEJ reporter by thinking through the design choices that were made during the construction of this tool.

  1. You will start by taking a close look at the vector that was selected. The product information for pMAX can be found here.
    • What features within the vector are involved in propagating the plasmid in bacterial cells? Why is propagation in bacteria useful?
    • What features within the vector are involved in expressing the plasmid in mammalian cells? What is the purpose of each?
  2. Compare the plasmid map of the pMAX-EGFP-MCS plasmid you will use to the parental pMAX vector.
    • What additions/deletions were performed to build pMAX-EGFP-MCS?
      Plasmid map of pMAX-EGFP-MCS.
  3. Now you will look more closely at the MCS regions.
    • The NHEJ reporter contains two MCSs. Remember an MCS (multiple cloning site) is a generic term for a collection of restriction enzyme recognition sequences.
    • Copy and paste the MCS1, nonsense insert, MCS2 sequence from the pMAX-EGFP-MCS plasmid into APE.
      • Highlight the following features: 1-32 bp = MCS1, 33-632 bp = nonsense insert, and 633-664 bp = MCS2.
    • Examine the sequence information for MCS1 and MCS2. What do you notice?
  4. Is either the MCS1 or MCS2 sequence in your NHEJ reporter the same as the MCS sequence in pMAX?
    • Use the APE file generated in Step #3 to generate a restriction enzyme map of the MCS1, nonsense, MCS2 sequence.
      • Go to EnzymesGraphic Map + U. What does the U denote?
    • Compare the restriction enzyme map for MCS1 and MCS2 to the depiction of the MCS in pMAX (found in the product information for pMAX linked above).
  5. Indeed the MCS from the pMAX parental vector was deleted and a new MCS was engineered for the pMAX-EGFP-MCS NHEJ reporter plasmid. Here are some considerations that went into this design choice:
    • Enzymes that are active in compatible buffers.
    • Enzymes that are on hand in the laboratory stocks.
    • Enzymes that are absent from pMAX and EGFP.
  6. Schematic of DNA damage substrates for NHEJ assay. For blunt ends one restriction enzyme is used. For compatible and incompatible overhangs two restriction enzymes are used (noted by the different colored DNA strands).
    But why include two MCS features that flank a nonsense insert? To answer this, consider the following details of the NHEJ assay. The pMAX-EGFP-MCS plasmid is both a substrate for and reporter of NHEJ in your experiment. First, the plasmid DNA will be damaged using restriction enzymes. The damaged DNA is then transfected into the M059K and M059J cells. Also, the presence of damaged DNA initiates NHEJ. If the damaged DNA is repaired via NHEJ, the pMAX-EGFP-MCS plasmid is restored and the CMV promoter drives transcription of the gene encoding EGFP and the cell will fluoresce green. Your goal for this aspect of the module is to assess the frequency of NHEJ when different types of DNA damage are used as the substrate. Specifically, as a class you will complete experiments with DNA that is damaged to have blunt ends, compatible overhangs, and incompatible overhangs.
    • So, back to our question...why were two MCS features included?
    • Hint: consider the benefits of being able to use two restriction enzymes in a double digest to generate damaged DNA.
  7. Next, consider the reason for the nonsense insert. In your Systems engineering research article you will use the class data to assess and compare NHEJ frequencies for all of the damaged DNA types. Given this, it is important that you know your DNA is damaged and that only the damaged DNA is transfected into the cells.
    • Why was the nonsense insert sequence included in pMAX-EGFP-MCS?
    • Hint: Review the M2D3 page and consider the size of the nonsense insert fragment.
  8. Lastly, why was the gene encoding EGFP included?
  9. Consider the damage types that will result from the digests below. Draw the DNA ends for each option.
    • PmeI
    • BglII and EcoRI
    • BglII and PstI-HF
  10. Before you leave, discuss with your partner which DNA damage type you will use in your NHEJ assay. Formulate a hypothesis that states whether the DNA damage you chose will result in a higher or lower frequency of NHEJ repair when compared to the other options. Include an explanation as to why you think you will see a higher or lower NHEJ frequency.
    • Sign-up for the DNA damage type on the Discussion page. Note: Selection is first-come-first-served!

Reagent list

From Boston Bioproducts:

  • RIPA Lysis Buffer
    • 50 mM Tris-HCl, pH 7.4
    • 150 mM NaCl
    • 1% NP-40
    • 0.5% sodium deoxycholate
    • 0.1% SDS
  • 100X Protease Inhibitor cocktail
    • AEBSF
    • Aprotinin
    • E-64 Besstain Leupeptin
    • EDTA
  • 6x Reducing Laemmli Sample Buffer
    • 375 mM Tris HCl, pH 6.8
    • 9% SDS
    • 50% glycerol
    • 9% betamercaptoethanol
    • 0.03% bromophenol blue

Precision Red Advanced Protein Assay from Cytoskeleton, Inc.

From Bio-Rad:

  • 4-20% Mini-PROTEAN TGX gel
  • TGS Buffer (25 mM Tris, 192 mM glycine, 0.1% (w/v) SDS, pH 8.3)
  • HiMark Pre-stained Protein Standard
  • Transfer Buffer (25 mM Tris, 192 mM glycine, 20% v/v methanol)

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