Biology 236: Plant Biology
Labs 5-7: Changing Protein Profiles During Maize Germination



Laboratory Objectives

  1. Demonstrate that protein profiles change during germination in maize.
  2. Determine whether the maize lea group 3 protein is found in pea embryos.
  3. Determine where in the maize seed the maize lea group 3 protein is localized via tissue printing.
  4. Acquaint you with the techniques of SDS-PAGE (Sodium Dodocyle Sulfonate-PolyAcrylamide Gel Electrophoresis) and Western analysis.


Polyacrylamide Gel Electrophoresis of Plant Proteins

Development is characterized by an ordered, controlled sequence of changes that occur within an organism at the molecular, cell, tissue and organ levels. Differentiation of cells and tissues involves processes that give rise to a variety of cell types within different tissues. Such cells within a particular tissue or organ are characterized by a unique morpholgical and chemical makeup.

Different parts of a seedling all contain different protein complements. Presumably, all cells within these different tissues have the same genetic constitution and, therefore, the same DNA encoded information in their chromosomes. Thus, if all cells contain the information necessary for the manufacture of all the different types of proteins found in the pea plant, what is it that controls the expression of this information? How does gene activity result in one protein complement in one part of the plant and an entirely different complement in another part? What controls the appearance of specific proteins in an organ at particular developmental stages? These are some major developmental questions that need answering.

The fact that different tissues within a plant contain unique complements of proteins was demonstrated by the above investigators, using the technique of polyacrylamide gel electrophoresis. In this technique a gel composed of polyacrylamide is polymerized between glass plates, across which is imposed an electrical potential that causes any appropriately charged macromolecule present to migrate through the gel. The preparation of these gels is a simple, rapid procedure in which the resulting pore size of the gel can easily be adjusted. The separation of the proteins is based solely upon their molecular size because the SDS acts to give all proteins a uniform charge.

As the proteins migrate through the large-pore gel (stacking gel), each protein species is concentrated at the boundary between the stacking and running gel in very thin disks (~25 µ), one on top of the other, in order of decreasing mobility. This steady-state stacking occurs within minutes after the voltage is applied, and one can observe this phenomenon by watching the concentration of the bromphenol blue sample dye into an extremely thin disk. Each protein is now free to migrate through the the smaller-pored running gel, each species separating from the others on the basis of their molecular sizes. After a sufficient time to achieve satisfactory separation of the protien disks in the running gel, the proteins are immobilized (fixed with methanol) and stained by placing the gels in coomassie blue. After destaining, the components of a particular protein extract can be visualized in the gel. To probe for specific antigens present in a complex protein sample, one could use specific primary antibodies for immunodetection (Western analyses). The specific antigen found by immune detection can then be compared to the electrophoretic pattern of the complete protein mixture. Molecular weight determinations are possible by including a lane of MW standards in the gel. See Fig. 1 for detail of enzymatic detection of membrane bound antigens, using the Enhanced Chemi-Luminescence (ECL) method of visualization.

Procedures

(see Fig. 1 for flow diagram of procedures)

Procedural Summary
  1. Week 1
  2. Week 2
  3. Week 3


Protein Extraction

NOTE: KEEP SAMPLES ON ICE AT ALL TIMES POSSIBLE TO PREVENT DEGRADATION OF PROTEINS.
  1. Label Eppendorf tubes for the various samples (e.g., maize zero time, 3 days after imbibition, 7 days imbibition, pea, and aleurone.).
  2. Scrape off the aleurone layer of zero time kernels (no other times) onto ice overlayed with aluminium. All groups combine your aleurone layers to make one sample.
  3. Separate maize embryos and seedlings from endosperm. I will demonstrate. Toss the endosperm.
  4. One sample at a time, mash the tissue in a mortar and pestle. Wipe clean the mortar and pestle between samples. Prepare the pea sample before the maize samples, and then prepare the maize from the most germinated to the least germinated.
  5. Add as little protein extraction buffer (PEB) as possible. Depending on the amount of tissue, add between 50 and 400 ul of PEB. [Alternatively, forego step 2 above, weigh the tissue to the nearest 0.00 mg, and then add 5X PEB. For example, if the sample weighs 100 µg, then add 500 µl of PEB].
  6. Grind the tissue to a uniform mash, put into labelled eppendorf tube, and vortex at high speed. The slurry should be able to spin when vortexed. If it does not move, add more PEB until it does.
  7. Put sample on ice, and then repeat steps 4-6 until all samples are processed.
  8. Heat the sample in a 65 degrees C bath for 5 minutes, vortexing once during that time.
  9. Vortex the sample again, hard and fast, for at least 2 minutes.
  10. Heat the samples in a 95 degrees C bath for 2 minutes.
  11. Vortex again for 1 minute.
  12. Cool for several minutes on ice.
  13. Spin in the microfuge for 5 minutes.
  14. Save top aqueous layer (save lower liquid phase along with pellet only if you have a marginal amount of sample).
  15. Repeat steps 11 and 12.
  16. Put your samples on ice.


Bradford Protein Micro Assay Procedure

(for protein quantification)
The Bradford Method is the most frequently used and reliable colorimetric method for determining protein concentration of a solution. The assay reagent utilizes an absorbance shift in the acidic coomaissie blue dye from 465-595 nm, when the dye binds to proteins. A comparison of the binding of the dye to an unknown protein to that of different amounts of a standard protein, usually bovine serum albumin (BSA) allows one to determine the approxiamte size of the specific bands.

Procedure
  1. Mix 1X Bradford Reagent (BR) bottle prior to use.
  2. Prepare a known protein concentration series by diluting a stock BSA (1 µg/µl) standard in the same diluent as the protein sample whose concentration is to be determined. The protein standard series should cover the range of concentration between 5 and 25 µg/mL. Convenient standard concentration data points are 0 (blank), 5, 10, 15, 20 and 25 µg/mL.
  3. Prepare 1X Bradford Reagent (if not already made). This is good for about 1 week. In duplicate, add 3 mL of 1X BR + 1-2 µl of protein sample. Be sure to prepare a blank of 1X BR.
  4. Vortex, then wait about 15 minutes.
  5. Determine the absorbance at 595 nm (A595) of samples on Spectrophotometer 20 after calibrating blank (Bradford reagent with no added BSA) to equal 0.000 Absorbance (100% Transmittance).
  6. Prepare a standard curve by plotting the absobance at 595 nm versus protein concentration. Use Cricket Graphics on the Mac to plot the data. Use the standard curve to determine the protein concentration for each unknown protein sample.

If the unknown protein concentraion is too high, dilute the protein, assay a smaller aliquiot of the unkown, or generate another standard curve in a higher concentration range (e.g., 10 to 100 µg).
If the samples are too dilute, then precipitate with Acetone + Beta-Mercaptoethanol.
  1. Add sample:acetone at ratio of 1:4 volumes.
  2. Leave in -20°C freezer for at least one hour.
  3. Quick spin in microfuge to pellet protein (do not spin long or else the pellet is difficult to get back into solution).


To make the tissue prints:

  1. With gloves on, obtain a 2x2 inch piece of Immobilon-P membrane.
  2. With a pencil, divide the membrane into 6 sections.
  3. Slice a maize kernel in half down the embryonic axis.
  4. Stick a tac through it.
  5. With the help of tweezers, press the cut edge of one half of the embryo. Repeat twice more. Toss the kernel half, and repeat with the second half on the bottom three spaces.
  6. Record the orientation of the embryo (sketch in your notebook).
  7. Label bottom right hand corner with your initials.
  8. Replace the protective blue paper to both sides of the membrane.
  9. Give to me, and I will store them in the fridge until the immunodetection lab.

  • SDS-PAGE


    NOTE: The SE 250 Mighty Small II is a miniature vertical slab gel unit intended for rapid electrophoresis of protein samples of small volume. Two gel sandwiches of polyacrylamide gels will be run simultaneously -- one for Coomassie staining for visualizing the changing protein profiles during development, a second gel for transfer a membrane for Western analysis. Polyacrylamide gels are run using precast gels (4% stacking and 15% running) sandwiched between plastic plates. The core of the unit serves as a heat exchanger through which coolant may be circulated.
    1. Attach the precast gel to the gel apparatus.
    2. Add 1x running buffer to lower and upper chambers [use about 500 mL of running buffer total].
    3. Prepare your samples to load. You will want to prepare enough for 30 µg of protein in each of 7 eppendorf tubes for good measure, but load only 25 µg total. Load 20 ug of protein in each lane for the gel to be stained, and then, with the same tip, load 5 ug of protein in the gel to be blotted.
      • Be sure to mark the gels with tape near the top of the gels.
      • Fill out worksheet, Fig. 2, in order to determine the appropriate volume of sample + buffer to load per well.
      • Load the same final volume in each lane (except standard lane). Use Type I water to make up the difference.
      • Load 5 µl of prestained standards in well 2 (second from left) on both gels.
    4. Load your samples from left to right (lanes 2 to 9), using the p-200 and long tips.
      • You will skip the outside lanes due to the "smiling" effect of heat distortion.
      • Use new tips for each different sample, but the same tip for duplicate lanes in the two gels.
      • Put the used tips in the dishpan provided.
    5. Run at ~200 Volts for one hour (or until dye front elutes from the gel).
      Be sure to stop the run before the orange marker runs off the gel.
    6. Turn the power off.
    7. Remove both gel sandwiches (plastic plate - gel - plastic plate) from the gel unit.
    8. Cut the upper right-hand corner so that you can properly orientate the gel when you view it after staining and de-staining it.
    9. Separate the gel to be stained from the plastic plates.
      Lift the front plate off with a spacer providing leverage from a corner. With the spacer, lift a corner of the gel.
    10. With two hands, transfer the gel to the tupperware of Coomassie Blue stain and shake gently on shaker for at least 30 minutes.
    11. Leave the gel to be used for Western analysis in the plastic plates.
      • Wrap in plastic wrap to keep it from drying.
      • On a piece of tap, label the gel with your initials.
      • Put the wrapped gel in the refridgerator for use in the next lab period.
    12. After the first gel is stained, move the gel to a tupperware with destain. Shake gently for 15-30 minutes until you clear the background.
      • If you leave it too long in the destain, even the proteins will become destained.
    13. Put your destained gel on a piece of saran wrap, grab its image on the ELMO, wrap the gel in the saran wrap, label it with you initials, and then put it in the refridgerator with your other gel.


    Misc. Info

    Prestained Markers

    Western Analyses of LEA Proteins

    Procedure for blotting with the Hoefer TE 70 SemiPhor.

    1. Soak your unstained gel in Western Transfer Buffer for ~ 10 minutes.
    2. Cut Whatmann 3MM filter paper and Immobilon-P membrane to the same length and width as the gel. You will need 2 sets of Whatmann 3 MM paper, 9 sheets in step 6.
    3. With gloved hands, immerse immobilon-P into a tupperware container containing 100% methanol for 1-2 seconds (the membrane is hydrophobic).
    4. Use tweezers to move the membrane to a tupperware container with Type I water. Shake it around a bit to dilute the methanol, which otherwise hampers protein transfer.
    5. Soak 3MM filter paper and Immobilon-P membrane in transfer buffer 15-20 minutes. Soak the filter paper as two units of 9 sheets. Put one end in the transfer buffer. As the liquid soaks into the paper, slowly lower the stack until all is soaked (this prevents pockets of air from forming, preventing transfer).
    6. Set up the gel unit as follows (see Fig. 3):

      transfer unit:
      Bottom: lower graphite electrode (anode) bottom
      9 sheets of filter paper (or 4 sheets of thicker paper)
      Immobilon-P membrane with cut in upper left corner
      gel with a cut in upper left corner for orientation later
      9 sheets of filter paper (or 4 sheets of thicker paper)
      Top:upper graphite electrode (cathode) top

    7. Roll a glass rod over the stack to force out air bubbles. Blot away excess buffer with Kim wipes.
    8. Put cathode cover of top of the stack, and be sure to plug in the wire from the cathode to the bottom unit. Plug the red and black electrodes into a power source (be sure that the power is OFF, the toggel switch is on low and that the other toggel switch is set on amperage).
    9. Transfer the proteins from the gel to the membrane at 0.9 mA/cm2 constant current for 1 hour. Determine the total surface area of all blots being transferred at one time.
    10. After the run is over, the Immobilon-P goes into blocking solution (Blotto). The gel should be stained in Coomassie blue to check for completeness of the transfer (higher molecular weight proteins are usually not transferred completely).


    Immunodetection

    NOTE: See Fig. 4 for schematic of ECL detection.
    1. After electrophoresis and blotting, block the membrane in BLOTTO solution for 30 minutes on shaker in seal-a-meal, or leave overnight at 4°C in cooler.
    2. Briefly rinse the membrane, using two changes of PBS-T, then wash once for 15 minutes, and twice for 5 minutes with fresh changes of the buffer at room temperature. Use as large a volume of PBS-T as possible.
    3. During the washing step, dilute the primary antibody.
    4. Transfer the membrane to a seal-a-meal, add primary antibody and agitate on nutator for at least one hour (could go overnight in 4 C cooler).
    5. Wash membrane as detailed in step 2.
    6. During the washing step, dilute the secondary antibody.
    7. Transfer membrane to seal-a-meal and incubate with secondary antibody on shaker for 30 minutes.
    8. Wash the membrane 1 X 15 minutes and 4 X 5 minutes in fresh changes of wash buffer.
    9. During the wash, prepare detection reagents 1 and 2 by mixing equal amounts of each into a 15 mL Falcon tube (need only enough to cover the membrane). Let warm to room temperature.
      Perform the following procedures in the dark room.
    10. Incubate membrane in detection reagents for 1 minute maximum.
    11. Drain off excess detection reagent by holding it on end, blot on Whatman 3MM filter paper, and wrap blots in plastic wrap (DO NOT allow blot to dry). Gently smooth out air pockets.
    12. Switch white lights off but leave red safe lights on.
    13. Cut off the upper RIGHT-hand corner of a piece of XOMAT film and then place the film in a film cassette.
    14. Place the blot, protein side down (so that the cut corner of the blot is in the same position as the cut corner of the film), in the upper left-hand corner of the cassette. Work as quickly as possible; minimize the delay between incubating the blot in detection reagent and exposing them to the film.
    15. Close the cassette and expose for 5 seconds to 5 minutes. (If bands of green light are visible in the dark, then opt for the shorter time). DO NOT move the film once exposed to membrane!
    16. Repeat steps 14 and 15 at different places on the film to provide various exposure times (you should be able to make 8 different exposures -- two across and four down).
    17. Develop your film in the Fisher Model K-Plus Automatic X-ray Film Processor.
      Start Up
      • Remove the top cover.
      • Remove the first metal plate.
      • Use hemostat to put plug in water chamber (rubber O-ring down).
      • Replace metal plate and top lid.
      • Turn on cold water (1/2 turn).
      • Turn machine on.
      • Wait 15-20 min for water tank to fill up (you will know that it is full when water comes out of the large hose in the sink).
      • The developer is ready when the Dev-at-temp light goes on.
      • Feed film through lengthwise when the Feed Film light goes on.
      • Can send through second film when Feed Film light goes on again. Start to finish takes 3 min. If it takes much longer, then your film has gotten stuck.
      • Remove the plastic lid. Remove the two metal covers, and fish you film out of one of the tanks or rollers.

      Shut Down
      • Turn machine off.
      • Turn water off.
      • Pull out plug for water tank.
      • Prop open the lid with water tank plug.
    18. Label the film with your initials.
    19. Capture image on ELMO.


    References

    Thomann EB, Sollinger J, White C and Rivin CJ (1992) Accumulation of Group 3 Late Embryogenesis in Zea mays Embryos. Plant Physiology 99, 607-614.


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