Peptide Design

The crucial first step in custom antibody development is peptide design. The specificity and utility of the antibody depends critically on the immunizing peptide. As an example of the peptide design process, consider figure below, illustrating the phospho-peptide for serine-40 in tyrosine hydroxylase. The sequence N-terminal and C-terminal to the phospho-serine is shown at the middle of the figure, while the peptide design is shown at the bottom.

The rationale for the design of this peptide is as follows:

Truncated N-Terminal sequence: First, the peptide has a truncated N-terminal sequence. This was done to eliminate cross reactivity with homologous proteins that were identified in a Blast search.

Use of a Short Sequence: Second, another feature of the peptide design was the use of a very short sequence which forces the phosphoseryl residue into the epitope recognized by the antibody.

Amidation of the C-Terminus: Third, given the very favorable amphipathic character of the peptide, the C-terminus was amidated to better emulate the native tyrosine hydroxylase protein. Such modifications can improve immunoreactivity in western blots and especially in immunohistochemical applications.

Addition of an N-Terminal Cysteinyl Residue: Lastly, an N-terminal cysteinyl residue was added for conjugation to the carrier protein and coupling to the affinity column.

Antibody Purification

Three antibody possibilities: After immunization with the phospho peptide, three types of antibodies can be produced in the rabbit’s immune response. As seen in Figure 1 below, the first is the phospho-specific antibody that is desired. However, antibodies that are specific for the dephospho form of the peptide can also be generated if phosphatases in the rabbit dephosphorylate the peptide conjugate that had been injected. Lastly, pan-specific antibodies can be generated against sequences/conformations of the peptide that do not involve the phosphoryl group in the epitope. These antibodies react with the protein irrespective of its phosphorylation state. The only way to isolate the desired phospho-specific antibody is through sequential phospho- and dephosphoaffinity chromatography.

Isolation of the Phospho-Specific Antibody: The first column we use is the phosphopeptide affinity column (see Figure 2). We first prepare an IgG fraction from the serum and apply it to our phosphopeptide affinity column. Two of the three types of antibodies that we described previously will bind to this column. First, the phospho-specific antibody (blue) will bind because the phosphopeptide is present on this column. The pan-specific antibody (red) will also bind because the sequence of the peptide that it recognizes is also present on this column. The one antibody that will not bind to this column is the antibody that is specific for the dephospho form of the peptide (green), as this form of the peptide is not present on the column. These latter antibodies pass through the column in the flow-through, together with the entire complement of additional IgGs that were isolated from the rabbit serum. These “flow-through” antibodies are saved, and the phospho- and pan-specific antibodies are then eluted from the phosphopeptide affinity column.

Characterization in Western and IHC

Specificity in Western Blotting

The composite Western blot below in Figure 1 shows the characteristic doublet of our synapsin pan antibody as well as three of our phospho synapsin antibodies, and demonstrates two very import aspects of antibody specificity. First, there are no cross-reacting bands labeled by the antibody in the rat brain homogenate. Second, phospho-specificity is definitively demonstrated by comparing the signal in untreated rat brain homogenate (left) to the same homogenate, but that has been treated with lambda and alkaline phosphatase prior to being run on the gel (right). Treatment with phosphatase had no effect on the pan antibody’s signal, but completely eliminated the immunolabeling of the phospho antibodies.

Immunohistochemistry

Now we turn to immunohistochemistry. Many of our antibodies have been tested in IHC. Figure 2 below shows staining of cultured neurons with anti-synapsin pan antibody in green (left), and of C57 mouse striatal cells with anti-synapsin Ser⁵⁴⁹ (right).

Figure 3 below illustrates our pan-TH antibody’s extensive labeling in this photomicrograph of the retina (left). In contrast, a phospho-TH antibody selectively labels only the two amacrine cells in this light-stimulated retina example (right).

Use a lysis buffer containing 1% SDS when preparing whole cell lysates to ensure membrane and other hard to solubilize proteins, even synaptic junction proteins, are completely solubilized.

• 1% SDS lyses everything, preserving proteins from protease and phosphatase enzymes. Additional cocktail inhibitors are not necessary when 1% SDS is used.

When preparing lysate samples use fresh BME in 4X sample buffer to ensure samples are properly reduced and denatured.

• BME reduces the sulfhydryl groups in proteins and is highly volatile. We recommend adding fresh BME to 4X sample buffer just prior to adding to your samples.

Stain your membrane with Ponceau S after transfer to confirm successful protein transfer and to determine the exact alignment of lanes and placement of the proteins on the gel.

• Ponceau S will show you where protein has/ has not transferred onto the membrane and allow you to visualize any areas where bubbles or other factors may have affected the transfer of protein from the gel to the membrane.
  
• Make sure to collect excess Ponceau S for reuse and to thoroughly rinse membranes in dH₂O after staining to remove any remaining excess Ponceau S. Ponceau S staining is reversible and will not interfere with antibody labeling of the membrane.

When testing multiple antibodies using the same lysate, try using a stacking layer with one large trough instead of multiple lanes to maximize the number of strips available for testing.

• Once the gel has been transferred to a blot, you can use a razor blade and ruler to cut the blot into strips of equal widths. In most cases, twice as many strips can be cut from one large trough blot than can be cut from a blot with multiple lanes. The strips can then be incubated in a tray with narrow wells.

Proteins of different molecular weights transfer from gel to membrane at different speeds and efficiencies. Proteins with larger molecular weights will take longer to transfer than proteins with lower molecular weights. Try staining your gel with coomassie after the transfer to confirm that the proteins of interest have migrated out of the gel. Remember to also consider protein loss across the membrane. By adding a second membrane behind the first, you can see to what extent the proteins have blown through. Process both membranes with your antibody. Use data from such experiments to optimize transfer time for the specific protein of interest.

When working with phospho-tyrosine antibodies, try blocking and incubating with 3% and 1% BSA respectively instead of milk. Milk often contains high concentration of phospho-tyrosine and thus can give an abnormally high background if used as a blocking agent with phospho-tyrosine antibodies.

Optimal primary antibody dilutions should be determined experimentally using a dilution curve. A dilution that is too low will give a signal that is oversaturated. A dilution that is too high will give a signal below the detection threshold. Finding the “Goldilocks” zone where the signal is detectable, but not saturated is the goal.

• Experimental protocols and imaging systems vary from lab to lab. Trying a dilution curve with the primary antibody allows you to determine the best dilution to use in your system.

Antibodies are sensitive and must be handled and stored with care to ensure they remain active and efficient in binding their target proteins. If you don’t plan on using all of the antibody right away, freeze it in aliquots (volume dependent on application) at -20C or lower when they are not being used and keep your “working aliquot” stored at 4C.

• Aliquots from antibodies formulated with 50% glycerol in their buffer ­can be repeatedly taken directly from the stock vial stored at -20C without freeze/thawing due to the glycerol content.

PhosphoSolutions’ ultimate guide to doing western blots is broken down into 5 sections, containing a detailed list of steps, buffers, and specific materials needed within each section. Additionally, helpful technical tips are offered throughout the protocol to provide insight about various factors that should be considered when performing western blotting.  Our main goal is to help researchers obtain publishable and reproducible results.

Let’s Begin…. Happy Blotting!

Sample Preparation:

Before you Begin:

• Prepare sample: 
  Lysate: Thaw if frozen, then sonicate lysate in recommended lysis buffer to break up any residual debris and for easy pipetting without clogging.
  Organs and Cells: Refer to ‘Lysate Preparation: Organs and Tissue Culture Cells’ protocol
• Heat Block: set to 95˚C

Materials Required:

• 4X Sample Buffer: 0.125 M TRIS, 8%(w/v) SDS, 40%(v/v) Glycerol, 20%(v/v) β-Mercaptoethanol (BME), pH 6.8 w/HCl
• Screw cap microfuge tubes
• Lysis Buffer: 1%(w/v) SDS, 10 mM TRIS, 1 mM EDTA, pH 8.0
• Vortexer
• Mini-centrifuge

1. Prepare lysate samples in screw cap microfuge tubes with appropriate amount of 4X sample buffer.
   Tech Tips:
   • Use fresh BME in 4X sample buffer to ensure samples are properly reduced and denatured.
   • Mix additional 4X sample buffer with lysis buffer to adjust final volumes for protein samples and to fill empty lane(s). Adjust the volume of the molecular weight marker so each lane runs proportional down the gel.
2. Vortex samples and boil for 5 minutes at 95^oC.
3. Allow samples to cool. Then vortex and spin down samples in mini-centrifuge to collect all the lysate before loading onto gel.

SDS PAGE:

Materials Required:

• Lysate sample
• Lysis Buffer: 1%(w/v) SDS, 10 mM TRIS, 1 mM EDTA, pH 8.0
• 4X Sample Buffer: 0.125 M TRIS, 8%(w/v) SDS, 40%(v/v) Glycerol, 20%(v/v) β-Mercaptoethanol (BME), pH 6.8 w/HCl
• 1X Running Buffer: 25 mM TRIS, 80 mM Glycine, 35 mM SDS, do not adjust pH
• Electrophoresis Apparatus and power source
• Gel loading pipette tips
• Protein Marker
• Deionized Water (dH₂O)
  
• 4X Lower Gel Buffer: 1.5 M TRIS, pH 8.8 w/ HCl
• 4X Upper Gel Buffer: 0.5 M TRIS, pH 6.8 w/ HCl
• 10% APS: 10%(w/v) Ammonium Per Sulfate dissolved in dH₂O
• 10% SDS: 10%(w/v) Sodium Dodecyl Sulfate dissolved in dH₂O
• 30% Acrylamide/Bis Solution (37.5:1)
• TEMED: N,N,N’,N’ – tetramethylethylene diamine
  

  ---

1. Prepare appropriate percentage SDS gel based on the molecular weight of the protein of interest. (Gel recipes and recommendations are listed below).
   Tech Tips:
   • Add TEMED and 10% APS as last ingredients of gel recipe since they are responsible for starting the polymerization process.
   • When testing multiple antibodies using the same lysate, try using a stacking layer with one large trough instead of multiple lanes to maximize the number of strips available for testing.
2. Assemble gels in the electrophoresis apparatus and add 1X running buffer to recommended fill level.
3. Using gel loading pipette tips, load cooled samples and protein marker onto gel. For a multi-lane gel load ~5-50 µg of protein per lane. For trough gel load ~0.5-1.0 mg of protein.
   Tech Tip:
   • When using multi-lane gels, load equal volumes of sample to each lane to prevent lateral band spreading. In any empty lanes, load a ‘blank buffer’ consisting of lysis buffer containing 4X sample buffer at the same volume as samples. Molecular weight ladder also must be brought up to same volume with the ‘blank buffer’ in multi-lane gels.
4. Run gels per electrophoresis apparatus’ manufacturer’s recommendations.
   Tech Tips:
   • Allow gel to run until the dye front has passed through the gel. The run time for the gel will vary depending on the percentage of the gel.  A 7.5% gel will run the fastest, a 15% the slowest.  Check gel frequently.
   • If gel front begins to exhibit a “smile”, reduce voltage.

Protein Transfer:

Before you begin:

• Prepare SDS PAGE gel: Soak gel in transfer buffer for 10 minutes to equilibrate after removing from electrophoresis apparatus.
• Cut Whatman filter paper: Cut 2 pieces of filter paper slightly larger than membrane to keep gel and membrane held together and to prevent any separation or slippage.

Materials Required:

• Transfer Buffer: 25 mM TRIS, 190 mM Glycine, 10% Methanol, do not adjust pH
• Transfer apparatus and power source
• 100% Methanol
• Membrane: PVDF or Nitrocellulose
• Coomassie Blue: 0.1%(m/v) Brilliant Blue R- 250, 40% Methanol, 10% Acetic Acid, 50% dH₂O
• Gel Destain: 10% Acetic Acid, 90% dH2O
• Ponceau S stain: 0.2% Ponceau S stain, 3% Trichloroacetic Acid, 3% Sulfosalicylic Acid, 94% dH₂O
• Plastic bag

1. Select either polyvinylidene difluoride (PVDF) or nitrocellulose membrane as the solid support for protein transfer. If using PVDF be sure to activate the membrane by soaking it in methanol for 1 minute prior to incubating it in transfer buffer for at least 15 minutes.
2. Assemble transfer ‘sandwich’ system sequentially based on manufacturer’s recommendation.
   
   Tech Tip:
   • Carefully remove any air bubbles present between the gel and the membrane with a roller, as bubbles will prevent proteins from transferring from the gel to the membrane.
3. Set the voltage, amperage, and time based on manufacturer’s recommendations.
   Tech Tip:
   • Larger molecular weight (~150-250 kDa) proteins will take a longer time to transfer than smaller molecular weight (~25-100 kDa) proteins. Adjust transfer times based on manufacturer’s recommendations and place an additional membrane in transfer to collect any protein that has potentially blown through the primary membrane.
4. After transfer, rinse membrane thoroughly in dH₂O to remove any remaining transfer buffer on the membrane and air dry the membrane until completely deactivated (solid white).
   Tech Tips:
   • After transfer, rinse gel with dH₂O and incubate gel in Coomassie Blue to verify transfer efficiency. Gently rock and soak gel for 30 minutes. Discard Coomassie Blue and soak in gel destain overnight rocking at room temperature. A Kimwipe can be placed in the corner of the box to speed up the destaining process.
   • Stain your membrane with Ponceau S after transfer to confirm successful protein transfer and to determine the exact alignment of lanes and placement of the proteins on the gel. Collect excess Ponceau S for reuse and make sure to thoroughly rinse membranes in dH₂O after staining to remove any remaining excess Ponceau S on the membrane and so protein bands are visible. Ponceau S staining is reversible and will not interfere with antibody labeling of the membrane.
5. Mark molecular weight markers and lanes on the membrane using a gel pen. The markers will fade over time or wash away in any buffer. For long term storage, seal protein blot in a plastic bag and store in dark or dimly lit area.

Immunolabeling:

Before you begin:

• Prepare membrane: Label and cut (if needed) membrane for immunolabeling.
  PVDF membrane: activate with 100% methanol for 30 seconds, and then rinse with dH₂O.
  Nitrocellulose: DO NOT SOAK NITROCELLULOSE MEMBRANE IN METHANOL.

Materials required:

• Wash Buffer (1X TTBS): 14 mM NaCl, 2 mM TRIS, 0.1%(w/v) Tween 20, pH 7.6
• Blocking Buffer: 5%(w/v) Non-Fat Dry Milk (NFDM) or 3%(w/v) Bovine Serum Albumin (BSA) in 1X TTBS
• Incubation Buffer: 1%(w/v) NFDM or 1%(w/v) BSA in 1X TTBS
• Primary Antibody
• Secondary Antibody: There are a variety of detection methods used, though antibodies conjugated with HRP are the most common and will be described in this protocol.
• 100% Methanol

1. Cut and label desired number lanes from membrane. If using a large trough gel for a single lysate, cut and label individual strips.
2. If using PVDF, membrane must be activated by immersing it in methanol for 30 seconds.
3. Block membrane in 5% nonfat dried milk (NFDM) in 1X TTBS for 30 minutes while rocking at room temperature to saturate any free binding sites on the membrane.
   Tech Tip:
   • If experiencing significant background noise, try using 3% BSA in 1X TTBS as the blocking buffer.
4. Using the manufacturer’s recommended dilution, dilute primary antibody in 1% NFDM in 1X TTBS, making sure final volume will completely cover membrane during incubation.
   Tech Tips:
   • Optimal primary antibody dilutions should be determined experimentally using a dilution curve.
   • If experiencing significant background noise, try using 1% BSA in 1X TTBS to dilute primary antibody.
5. Incubate membrane in primary antibody overnight while rocking at 4^oC.
6. Discard primary antibody solution and wash membrane using 1X TTBS 3 x 5 minutes at room temperature.
7. Incubate membrane in HRP conjugated secondary antibody at a dilution of 1:10,000-1:30,000 in 1% NFDM in 1X TTBS while rocking for 1 hour at room temperature.
   *Make sure to select appropriate secondary antibody depending on the primary antibody’s host species.
8. Discard secondary antibody solution and wash membrane using 1X TTBS 3 x 5 minutes.
   Tech Tip:
   • When working with a new antibody it is recommended that both BSA and NFDM be tested to optimize signal strength and quality.

Imaging and Data Analysis

Before you begin:

• Prepare imaging system: Warm up camera in machine and open software program for imaging.
  OR
• Dark room and film: Warm up dark light and layout all equipment used to easily find in minimal light.

Materials Required:

• Imagining system or film and dark room
• ECL detection substrate
• Conical tube- for mixing ECL substrate

1. Prepare chemiluminescent imaging system. If using film, prepare film and box in dark room.
2. Using the manufacturer’s recommended instructions, prepare ECL detection substrate in 15 mL conical tube. Determine optimal volume to cover membrane completely.
3. Incubate membrane in substrate for time indicated by ECL manufacturer. (Time may vary depending on sensitivity of substrate used).
4. ECL detect with imaging system or with film.
   Tech Tip:
   • If testing an antibody for the first time, it is recommended that multiple exposures are captured at varying lengths of time to optimize signal quality.
5. Quantitate western blot data using imaging system’s software or a preferred standalone software if available.

∗ Multiple bands within a western blot raise a critical flag concerning an antibody’s specificity. Any antibody that produces multiple banding in western blot should not be used in IHC unless additional testing can be performed to validate the specificity of the antibody.

PhosphoSolutions’ step-by-step protocol shows how to properly lyse organs and tissue culture cells when focusing on preserving protein phosphorylation, including detailed differences in lysing adherent and suspension tissue culture cells.  Within each section there is a detailed list of steps, buffers, and specific materials to help researchers through this process. Additionally, helpful technical tips are offered throughout the protocol to provide insight about various factors that should be considered when lysing organs and tissue culture cells. Our main goal is to provide essential techniques or tools to help researchers obtain publishable and reproducible results.

Before you Begin:

• Determine Protein Mass:
  Organs: Weigh fresh or thawed organs. If frozen, brush off frozen debris from organ. If fresh, wash organ with 1X PBS and dry with Kimwipe. Estimate protein mass as 10% of total mass.
  TC Cells: Make estimate based on confluency (adherent) and cell count (suspension).
• Heat Block: Set to 95^oC.
• Sonicator: Select probe and optimize strength based on lysis buffer volume and tube size to avoid foaming.

Materials Required:

• Lysis Buffer: 1%(w/v) SDS, 10 mM TRIS, 1 mM EDTA, pH 8.0
  *room temperature for organs
  *95˚C for adherent and suspension TC cells
• 1X PBS: 137 mM NaCl, 28 mM Na₂HPO₄, 5.4 mM KCl, 2.9 mM KH₂PO₄, pH 7.6
• Cell Scraper: For adherent tissue culture cells
• Dissection tools, scalpel, spatula, scissors
• Plastic transfer pipette
• Conical tube or microtube with screwcap
• Ice/water

Prepare Samples

A. Organs – Whole cell lysate

1. Place organ in conical tube or microfuge tube with screw cap. For large organs cut into 1/4” sections. For organs that are thick and dense cut even smaller sections.
2. Add lysis buffer prepared at room temperature. Choose a volume based upon desired concentration, which is based on the estimated mass. Keep sample at room temperature.
   Tech Tips:
   • 1% SDS lyses everything, preserving proteins from protease and phosphatase enzymes. Cocktail inhibitors are not necessary.
   • A concentration below 10 mg/ml is recommended for optimal lysing. Calculate concentration with formula below:
     

B. Adherent Tissue Culture Cells

1. Wash cells with 2-10 mls of room temperature 1X PBS once. Remove wash with transfer pipet.
2. Add 95^oC lysis buffer, enough to cover the entire plate.
   Tech Tips:
   • Recommended volumes: 2 mls for 150 mm plate, 1 ml for 100 mm plate, 500 ul for 60 mm plate.
3. Rock and rotate the plate to thoroughly coat cells.
   Tech Tips:
   • Adherent cells will immediately lyse and a glob of cells in lysis buffer will be present.
4. Transfer to a conical tube or microfuge tube with screwcap.
   Tech Tips:
   • Tilt plate and gently guide lysed cells with cell scraper to collect at the bottom of the plate. Cut off the tip of a plastic transfer pipette to help collect and transfer the large glob of cells into conical tube.

C. Suspension TC Cells

1. Pipette media and cells into conical tube.
2. Pellet cells by centrifugation (1 minute at 1800 x g) and remove media.
3. Resuspend cells with 1-3 mls of room temperature 1X PBS to wash.
4. Repeat step #2.
5. Add 95^oC lysis buffer.

Lyse Samples

1. Sonicate sample in 5-20 second intervals until buffer is clear and can be easily pipetted without clogging.
   Tech Tips:
   For optimal lysing:
   • To prevent overheating, place conical or microfuge tube in an ice/water bath while sonicating. Do not keep sample in ice bath for an extended period of time.
   • To avoid puncturing/melting the plastic tube, minimize prolonged contact with the probe to the conical tube walls.
   • To avoid foaming, keep probe tip in lysis buffer until sonication is complete.
   • If the lysate sample clogs while pipetting, sonicate again until clog is no longer present.
2. Heat sample at 95^oC for 10 minutes. Cool to room temperature.
   Tech Tips:
   If lysate appears:
   • Cloudy: Add 5% by volume increments of lysis buffer to sample, then gently rock until sample is clear in solution. A protein concentration below 10 mg/ml is preferred. A concentration above this will appear cloudy and insufficiently lysed.
   • “Goopy”: Repeat sonication until sample is easily pipetted.
3. Centrifuge lysate at 1800 x g for 5 minutes to pellet cell debris.
4. Repeat steps 1-3 if debris is present. Place lysate in fresh conical or microfuge tube. Save/discard old tube and debris.
   Tech Tips:
   • Organ and cell membranes will appear as the white, stringy debris in lysate. It is essential to completely lyse the debris when studying transmembrane proteins.

Determine Protein Concentration

Determine protein concentration using BCA, Bradford, or Lowry assays.

Determining phosphospecificity of phosphospecific antibodies is serious business to PhosphoSolutions. This step-by-step method is broken down into 2 major sections on how to dephosphorylate proteins the right way. Within each section there is a detailed list of steps, buffers, specific materials, and calculation examples to help researchers through this comprehensive process. Additionally, helpful technical tips are offered throughout the protocol to provide insight about various factors that should be considered when performing protein dephosphorylation. Our main goal is to provide essential techniques and tools to help researchers obtain publishable and reproducible results.

Dephosphorylation of Proteins Fixed on a Membrane

Materials Required:

• Protein Membrane Blot:
  -PVDF membrane is recommended.
  -Refer to Western Blot Protocol (up to Protein Transfer) to prepare protein membrane blot.
• Lambda Phosphatase Enzyme: Sigma Aldrich product, includes 10X phosphatase buffer and 10X MnCl₂, Catalog number: P9614
• 1X Incubation Buffer: Lambda phosphatase buffer and MnCl₂ solution diluted to 1X with dH₂O
• Alkaline Phosphatase Enzyme: Sigma Aldrich product, Catalog number: P0114
• Conical tube or microtube with screwcap
• Deionized H₂O (dH₂O)
• Wash Buffer (1X TTBS): 14 mM NaCl, 2 mM TRIS, 1%(w/v) Tween 20, pH 7.6
• 100% Methanol

This protein dephosphorylation method uses a two-step approach to focus on specificity and potency. The first treatment is done with a lambda phosphatase to specifically target phosphate groups from threonine, serine, and tyrosine amino acids for dephosphorylation. Most phosphatases only dephosphorylate serine and threonine amino acids. If you are testing the phosphospecificity of a tyrosine phosphospecific antibody, lambda phosphatase is essential. The second dephosphorylation treatment is done with an alkaline phosphatase, a significantly more potent enzyme than lambda phosphatase. The alkaline phosphatase only dephosphorylates phosphate groups from threonine and serine amino acids, and therefore is not intended to be utilized alone to determine the phosphospecificity of tyrosine phosphospecific antibodies. When using both lambda and alkaline phosphatases in conjunction with each other, it is best to separate the treatments so the alkaline phosphatase doesn’t inhibit the activity of the lambda phosphatase. There are a variety of phosphatases one may use to dephosphorylate proteins; the ones listed in this protocol are recommended examples. The end user is highly encouraged to optimize their chosen phosphatase(s) before determining phosphospecificity of an antibody.

In this protocol, we refer to the phosphatase treated section of membrane (Treated) and a non-treated section of membrane (Control).

Membrane Preparation

A. Freshly Prepared Membrane

1. After protein transfer to PVDF is complete, rinse the membrane with dH₂O.
2. Cut membrane and place each portion into a separate container containing dH₂O.
   Tech Tips:
   • It is ideal to use a membrane prepared from a large one trough gel. Cut the membrane in half for each section.
   • If using multiple lanes, identify the lanes by staining the membrane with Ponceau S. Cut the fresh, wet membrane for each treatment. Place membrane sections back into container with water. Completely remove Ponceau S stain with 3 x 5 minute washes with wash buffer. Additional washes may be needed.
   • To distinguish between the two sections and orientation, make a small cut in the top right corner of each membrane and label the containers Control and Treated.
   • When labeling membrane sections a gel pen is needed so the ink does not wash away. However, the membrane can only labeled once it is dry.
3. Rock the membrane in wash buffer for 5 minutes at room temperature. DO NOT DRY MEMBRANE BEFORE TREATMENT.
   Tech Tip:
   • Choose a container for each membrane section that minimizes the volume needed to completely immerse blot in wash buffer.
4. Proceed to ‘Protein Dephosphorylation’.

B. Deactivated Membrane

1. Cut and label dry deactivated membrane.
   Tech Tips:
   • When performing a single phosphospecificity test, strips can be cut and used instead of treating a large section of membrane. This is optimal for saving special, hard to prepare lysate.
   • When labeling membrane sections or strips a gel pen is needed so the ink does not wash away.
   • Membrane strips and sections can be labeled with “C” for Control and “T” for Treated before activating the membrane.
2. Activate the two halves of the PVDF membrane by immersing them in methanol for 30 seconds in selected containers, then thoroughly rinse the membrane halves with dH₂O.
   Tech Tip:
   • Choose a container for each membrane section that minimizes the volume needed to completely immerse blot in wash buffer.
3. Wash membranes in wash buffer for 3 x 5 minutes.
   Tech Tip:
   • If the membrane was stained with Ponceau S, additional washes may be needed. Stain must be completely washed away from membrane before proceeding.
4. Calculate the total volume of 1X incubation buffer needed to completely immerse all Control and Treated membrane sections or strips in each container.
5. Proceed to ‘Protein Dephosphorylation’.

Protein Dephosphorylation

1. Prepare the calculated total volume of 1X incubation buffer needed to completely immerse all membrane sections or strips. Divide the 1X incubation buffer into 2 fractions: one labeled Treated and one labeled Control.
2. Add 5 uL of lambda phosphatase per mL of incubation buffer to the containers labeled Treated.
3. Remove wash buffer from membrane and add prepared Control and Treated solutions to respective membrane sections or strips. Incubate for 4 hours rocking at room temperature, or overnight rocking at room temperature.
   Tech Tip:
   • A sealed container is recommended to prevent evaporation.
4. Add 1 uL of alkaline phosphatase per mL of incubation buffer to the containers labeled Treated. Incubate for an additional 30 minutes, rocking at room temperature.
5. Discard 1X incubation solution. Rinse membrane thoroughly with dH₂O.
   Tech Tip:
   • If needed, restain the membrane halves with Ponceau S to visualize proteins and remove excess membrane or cut lanes.
6. Deactivate membrane by rinsing with dH₂O and air dry. The membrane is ready for blocking and antibody incubation (part of Western Blot protocol) once it is completely deactivated and the proteins are fixed to the membrane.
   Tech Tip:
   • To speed up the drying process, rinse the membrane with 100% methanol after rinsing with dH₂O.

Dephosphorylation of Proteins in a Lysate

Materials Required:

• 10X Lysis Buffer: 100 mM TRIS, 100 mM NaCl, pH 8.0
• Detergent for Lysis Buffer: 10% (v/v) NP40-LB
• 10% SDS: Phosphatase deactivating agent
• Lambda Phosphatase Enzyme: Sigma Aldrich product that includes 10X Phosphatase Buffer and 10X MnCl₂, Catalog number: P9614
• Alkaline Phosphatase Enzyme: Sigma Aldrich product, Catalog number: P0114
• 1X Lysis/Incubation Buffer: Prepared from 10X Lysis buffer, 10X Phosphatase Buffer, and 10X MnCl₂ solution
• Spatula, plastic transfer pipette, and scissors
• Conical tube or microtube with screwcap
• Ice
• Deionized H₂O (dH₂O)
• Heat block: Set to 95°C
• Sonicator: Select probe and optimize strength based on lysis buffer volume and tube size to avoid foaming of the protein sample.

This protein dephosphorylation method uses a two-step approach to focus on specificity and potency. The first treatment is done with a lambda phosphatase to specifically target phosphate groups from threonine, serine, and tyrosine amino acids for dephosphorylation. Most phosphatases only dephosphorylate serine and threonine amino acids. If you are testing the phosphospecificity of a tyrosine phosphospecific antibody, lambda phosphatase is essential. The second dephosphorylation treatment is done with an alkaline phosphatase, a significantly more potent enzyme than lambda phosphatase. The alkaline phosphatase only dephosphorylates phosphate groups from threonine and serine amino acids, and therefore is not intended to be utilized alone to determine the phosphospecificity of tyrosine phosphospecific antibodies. When using both lambda and alkaline phosphatases in conjunction with each other, it is best to separate the treatments so the alkaline phosphatase doesn’t inhibit the activity of the lambda phosphatase. There are a variety of phosphatases one may use to dephosphorylate proteins; the ones listed in this protocol are recommended examples. The end user is highly encouraged to optimize their chosen phosphatase(s) before determining phosphospecificity of an antibody.
Calculations for the recommended examples can be found at the end of the protocol. It is important to know the specific activity of the phosphatase(s) chosen to determine the amount of enzyme needed. If dephosphorylating pure protein and the molecular weight is known, use this number to determine the amount of each phosphatase needed. If you are preparing a lysate with mixed proteins an assumption of the average molecular weight, aka highly educated guess, is best used. The calculated phosphatase volumes are a starting point to determine the amount needed to dephosphorylate the protein. Since the calculation is based on assumption, additional experimental optimization may be needed.

Lysate Preparation

1. Determine the protein mass of the organs or tissue culture cells.
   Tech Tips:
   • Organs: Brush off any debris from frozen organs. Wash fresh organs with 1X PBS and dry with Kimwipe. Weigh fresh or thawed organs. Estimate protein mass as 10% of total mass.
   • Suspension Cells: Make estimate based on cell count.
   • Adherent Cells: This protocol is not recommended for adherent cells due to non-optimal lysing conditions. The ‘Dephosphorylation of Fixed Protein on membrane’ protocol is recommended.
2. Based on the estimated protein mass for the samples, calculate a desired final concentration of lysate.
   Tech Tip:
   • A final concentration below 10 mg/mL is recommended for optimal lysing. Calculate concentration with formula below:
     
3. Determine the volume needed for the desired protein concentration and prepare 1X lysis/incubation buffer. Chill on ice for 30 minutes before lysing.
   Tech Tips:
   • Bring all lysis/incubation buffer components to a final concentration of 1% or 1X.
   • 10% NP40-LB or 10% Triton are recommended detergents to use in the preparation of 1X lysis buffer. These detergents preserve proteins and lyse most enzymes, but do not lyse phosphatase enzymes.
   • 1% SDS lyses everything, including phosphatase enzymes. It is important not to use 10% SDS prior to dephosphorylation treatment.
4. In an appropriate volume conical tube or microtube, add the calculated amount of freshly prepared chilled lysis buffer to organs/cells to reach the desired concentration. Keep sample chilled on ice. Make a note of the volume of lysis buffer added.
   Tech Tips:
   • Organs: For large organs, cut into 1/4” sections. For organs that are thick and dense, cut even smaller sections.
   • Suspension Cells: Pipette media and cells into conical tube. Pellet cells by centrifugation
     (1 minute at 1800 x g) and remove media. Resuspend cells with 1-3 mLs of room temperature 1X PBS to wash. Repeat centrifugation to pellet and remove 1X PBS. Add desired volume of lysis buffer.
5. Sonicate sample in 5-20 second intervals until solution is clear and can be easily pipetted without clogging. Keep sample on ice while sonicating and after sonication is completed.
   Tech Tips:
   • Organ and cell membranes will appear as white, stringy debris in lysate. It is essential to completely lyse this debris when studying transmembrane proteins.
   • To avoid puncturing/melting the plastic tube, minimize prolonged contact with the probe to the conical tube walls.
   • To avoid foaming, keep probe tip submerged in lysate until sonication is complete.
   • Pipette sample after sonicating. If the lysate clogs while pipetting, sonicate again until clog is no longer present.
6. Divide lysate into 2 equal volumes: Label one the control lysate, which will not be treated with phosphatase enzymes. Label the other the treated lysate, which will be treated with the phosphatase enzymes. Make a note of the volume for each sample.
7. Spike the control lysate with 10% SDS to a final concentration of 1% SDS. Make note of volume of 10% SDS added to the control lysate. Heat control lysate at 95˚C for 10 minutes. Set sample aside, leave at room temperature.

Protein Dephosphorylation

1. Add 1 ul of lambda phosphatase per 1 mg of protein to the treated lysate. Make a note of the volume of enzyme added to the treated lysate. Incubate in a water bath at room temperature for 30 minutes.
   Tech Tip:
   • Testing the dephosphorylated lysate is advised before using the lysate to determine phosphospecificity of an unknown phospho-antibody. Try using a known phosphospecific antibody that is phosphorylated at the same type of residue, i.e. Ser, Thr, or Tyr as your unknown. Since the original calculation was based on assumption further optimization of the amount of enzyme and incubation time may be required.
2. Add 1 ul of alkaline phosphatase per 10 mg of protein to the treated lysate. Make a note of the volume of enzyme added to the treated lysate. Incubate in a water bath at room temperature for 10 minutes.
3. Add 10% SDS to the treated lysate to a final concentration of 1% SDS and place in a heat block at 95˚C for 10 minutes to inactivate the phosphatase enzymes. Make note of volume of 10% SDS added to the treated lysate.
4. Add together the recorded volumes of reagents added to each lysate in previous steps. Adjust the lysates to equal volumes using the freshly prepared lysis buffer that contains 1% SDS.
5. Determine the protein concentration using the control lysate.
   Tech Tips:
   • The prepared 10X buffer for the enzyme used in this protocol contains DTT, this reagent will interfere with the BCA assay. It is recommended to use a different assay to determine protein concentration.
   • It is not recommended to use the treated lysate to determine the protein concentration as the concentration will be higher than the control due to the presence of the phosphatase enzyme.
6. Lysate is ready for assay application or can be frozen at -80˚C for long term storage.
   Tech Tip:
   • When producing a large preparation of treated cells or organs, aliquoting the lysates into small volumes for individual assay applications is recommended for convenience.

Phosphatase Calculation

Important factors, physical constants, and assumptions:

• Avogadro’s Number: 6.023 x 10²³ molecules/mol
• Assumed average molecular weight of the protein lysate: 100,000 daltons (g/mol)
• The Specific Activity of the Phosphatases
  Lambda Phosphatase: 1 Unit hydrolyzes 1 nmol of phosphate per minute; 1 ul of lambda phosphatase contains 400 units, so for every 1 ul of phosphatase 400 nmol of phosphate will be hydrolyzed per minute.
  Alkaline Phosphatase: 1 Unit hydrolyzes 1 mmol of phosphate per minute; 1 ul of alkaline phosphatase contains 133 units, so for every 1 ul of phosphatase 133 mmol of phosphate will be hydrolyzed per minute.
• If you know the exact size and amount of that pure protein, use those numbers. If not, don’t freak out! An assumption (aka highly educated guess), is best used in these scenarios. The phosphatase volumes calculated are starting points to determine the amount needed to dephosphorylate 10 mgs of protein. Since the calculation is based on assumption, additional experimental optimization may be needed.

Lysate Calculation Example

The first calculation is to convert the protein lysate sample from milligrams to molecules. This amount is important in determining the amount of each phosphatase needed. Since the lysate is a mixture of proteins an assumed average molecular weight of 100,000 daltons (g/mol) will be used to simplify the calculation. In this example there are 10 mgs of protein (2mLs of lysate at 5 mg/mL) that will be treated with phosphatase enzyme.

Average Protein Molecular Weight: 100,000 g/mol

There are ~6.023 x 10¹⁶ molecules for 10 mgs of protein.

Lambda Phosphatase Calculation

The second calculation will be converting the number of lambda phosphatase units into molecules/minute. It is important to note that the reaction time and volume of enzyme can be adjusted once this has been determined.

For every 1 ul of lambda phosphatase enzyme added to the reaction, it will hydrolyze 2.41 x 10¹⁷ phosphate molecules. This final calculation can now determine how long the reaction will take.

It takes 15 seconds to completely dephosphorylate 10 mg of protein when using 1 ul of lambda phosphatase.

Alkaline Phosphatase Calculation

The same steps will be done to calculate the amount of alkaline phosphatase needed for the reaction. Notice the specific activity of Alkaline Phosphatase is different than Lambda Phosphatase.

It takes 0.045 seconds to completely dephosphorylate 10 mg of protein using 1 ul of alkaline phosphatase.

by Amy Archuleta

Have you ever wondered exactly what is happening in an SDS PAGE system when you turn on the power source and the wires start bubbling?

You are not alone! Here are the answers to the science behind all those different pHs and gel layers.

SDS-PAGE Basics

What exactly is SDS-PAGE?
It is an acronym for Sodium Dodecyl Sulfate–Polyacrylamide Gel Electrophoresis.
SDS is a detergent, an anionic (negatively charged) surfactant (compound that lowers surface tension). In the case of proteins, SDS disrupts the non-covalent bonds in protein molecules.
PAGE is a biochemical technique that allows for proteins to be separated by their electrophorectic mobility (how fast they move in an electric field). In the case of SDS-PAGE, they are separated by their size (molar mass), and not their charge.

What causes the movement of the molecules through the gel?
An electric current. When you put the lid on your gel box and turn on the current, the negatively charged proteins will try to move through the gel towards the positively charged anode. The cathode and anode are the wires in your tank that are bubbling once you turn on the system.

What causes all those bubbles?
H₂ and O₂. Once the electric current is applied, the anode and cathode are involved in redox reactions that remove electrons from water molecules in the running buffer, resulting in gas formation. At the negatively charged cathode, positively charged hydrogen ions become hydrogen gas. At the positively charged anode, negatively charged oxygen ions become oxygen gas. You may observe more bubbles at the cathode than at the anode. This is because there are two hydrogen atoms for every one oxygen in a water molecule. There will be twice as many hydrogen gas molecules formed.

The Chemical Ingredients and What they Do

What exactly does SDS do?
It unfolds proteins. Application of SDS to proteins causes them to lose their higher order structures and become linear. Since SDS is anionic (negatively charged), it binds to all the positive charges on a protein, effectively coating the protein in negative charge.

Why do we want the protein coated in negative charges?
To remove charge as a factor in protein migration through the gel. SDS binds to proteins with high affinity and in high concentrations. This results in all proteins (regardless of size) having a similar net negative charge and a similar charge-to-mass ratio. In this way, when they start moving through a gel, the speed that they move will be dependent on their size, and not their charge.

After getting hit with SDS, is a protein’s size the only thing that affects its migration through the gel?
It is by far the biggest factor. However, SDS can bind differently to different proteins. Hydrophobic proteins may bind more SDS, and proteins with post-translational modifications such as phosphorylation and glycosylation may bind less SDS. These effects are usually negligible, but not always, and should be considered if your protein is running at a different molecular weight than expected.

What is in the running buffer?
Tris, glycine, and SDS, pH 8.3. Tris is the buffer used for most SDS-PAGE. Its pKa of 8.1 makes it an excellent buffer in the 7-9 pH range. This makes it a good choice for most biological systems. SDS in the buffer helps keep the proteins linear. Glycine is an amino acid whose charge state plays a big role in the stacking gel. More on that in a bit.

What is in the sample loading buffer?
Tris-HCl, SDS, glycerol, beta mercaptoethanol (BME), Bromophenol Blue. This is the buffer you mix with your protein samples prior to loading the gel. Again with the Tris buffer and its pKa. The SDS denatures and linearizes the proteins, coating them in negative charge. BME breaks up disulfide bonds in the proteins to help them enter the gel. Glycerol adds density to the sample, helping it drop to the bottom of the loading wells and to keep it from diffusing out of the well while the rest of the gel is loaded. Bromophenol Blue is a dye that helps visualization of the samples in the wells and their movement through the gel. Sample loading buffer is also known as Laemmli Buffer, named after the Swiss professor who invented it around 1970.

What is in the gels?
Tris-HCl, acrylamide, water, SDS, ammonium persulfate, and TEMED. Although the pH values are different, both the stacking and resolving layers of the gel contain these components. Tris and SDS are there for the reasons described above. Ammonium persulfate and TEMED work together to catalyze the polymerization of the acrylamide. The Cl- ions from the Tris-HCl work with the glycine ions in the stacking gel. Again, more to come on that.

Acrylamide Sets the Pace

What is in the gel that causes different sized protein molecules to move at different speeds?
Pore size. When polyacrylamide is combined in solution with TEMED and ammonium persulfate, it solidifies, effectively producing a web in the gel. It is through this web that the linearized proteins must move. When there is a higher percentage of acrylamide in the gel, there are smaller pores in the web. This makes it harder for the proteins to move through the gel. When there is a lower percentage, these pores are larger, and proteins can move through more easily.

Why are there different percentages of acrylamide in gels?
To optimize the resolution of different sized proteins. Different percentages of acrylamide change the size of the holes in the web of the gel. Larger proteins will be separated more easily in a gel that has a lower percentage of acrylamide – because the holes in the web are larger. The reverse is true for smaller proteins. They will resolve better in a gel with a higher acrylamide percentage because they will move more slowly through the holes. Small proteins will fly through a low percentage gel and may run off the end of the gel.

Gel Layers: It Takes Two

WHAT are there two layers in the gel?
The stacking layer and the resolving layer. The top (stacking) layer has a lower percentage of acrylamide and a lower pH (6.8) than the bottom (resolving) layer, which has more acrylamide and a higher pH (8.8). SDS PAGE is run in a discontinuous buffer system. There is discontinuity not only between the gels (different pH values and acrylamide amounts), but also between the running buffer and the gel buffers. The running buffer has different ions and a different pH than the gels.

WHY are there two layers in the gel?
They have different functions. The stacking layer is where you load your protein samples. The purpose of the stacking layer is to get all of the protein samples lined up so they can enter the resolving layer at exactly the same time. When you load a gel, the wells are around a centimeter deep. If your samples entered the resolving layer this spread out, all you would see is a big smear. The resolving layer then separates the proteins based on molecular weight.

How does the stacking layer do its job?
Low acrylamide content and low pH. The low percentage of acrylamide in the stacking layer allows for freer movement of the proteins and helps them line up to enter the resolving layer together. The lower pH allows glycine to be in its zwitterionic state.

Wait – did you just sneeze?
Close. I said glycine is a zwitterion at pH 6.8 in the stacking buffer.

It’s All About the Glycine

So what’s up with glycine?
A lot. It is the key to the discontinuous buffer system. It is the ionic state of glycine that really allows the stacking buffer to do its thing. Glycine is an amino acid with the chemical formula NH₂-CH₂-COOH. The charge of its ion is dependent on the pH of the solution that it is in. In acidic environments, a greater percentage of glycine molecules become positively charged. At a neutral pH of around 7, the ion is uncharged (a zwitterion), having both a positive charge and a negative charge. At higher pHs, glycine becomes more negatively charged.

What does glycine’s charge have to do with the stacking layer?
Everything. Glycine is in the running buffer, which is typically at a pH of 8.3. At this pH, glycine is predominately negatively charged, forming glycinate anions. When an electric field is applied, glycinate anions hit the pH 6.8 stacking buffer, and change to become mostly neutrally charged glycine zwitterions. That means they move slowly through the stacking layer toward the anode due to their lack of charge.

By contrast, the Cl- ions (from the Tris-HCl in the gel) move at a faster rate towards the anode. When the Cl- and glycine zwitterions hit the loading wells with your protein samples, they create a narrow but steep voltage gradient in between the highly mobile Cl- ion front (leading ions) and the slower moving, more neutral glycine zwitterion front (trailing ions). The electromobilities of the proteins in your sample are somewhere in between these two extremes, and so your proteins are concentrated into this zone and herded through the stacking gel between the Cl- and glycine zwitterion fronts.

What happens to glycine zwitterion in the resolving layer?
It gets real negative, real fast. When the Cl- and glycine zwitterion fronts hit the resolving layer at a pH of 8.8, the glycine ions gain a lot of negative charges. They are no longer predominately neutral and take off towards the positively charged anode as glycinate anions. Unaffected by polyacrylamide, they speed past the protein layer, depositing the proteins in a tight band at the top of the resolving layer.

Final Resolution

What happens to the proteins in the resolving layer?
They slow way down and start to separate. The proteins moved more easily through the stacking layer because of the low percentage of acrylamide. Now that they are starting into the resolving layer which has a higher percentage of acrylamide, they have to slow down. Also, without the voltage gradient from the Cl- and glycine zwitterion fronts, they can separate.

How does this all end?
Hopefully with beautifully tight bands separated by molecular weight. The different sized proteins run at different speeds through the gel, the big ones taking longer as they try to navigate the polyacrylamide web. The point at which they stop moving is dependent on when you turn off the power source. A good time to do this is usually when the dye-front running ahead of your protein samples (the blue line) reaches the very end of the gel. If you used the correct percentage of acrylamide, the molecular weight range of your protein of interest should be separated perfectly along the length of your gel!

The VERY Basics of Western Blotting

by Amy Archuleta

What is Western Blotting?

The Western Blot (or immunoblot) technique uses antibodies to detect protein targets that have been bound to a membrane. It was introduced in 1979 by Harry Towbin’s research lab in Switzerland.

Why is it called “Western”?

“Western” is a play on words based on a similar technique. Edwin Southern published a DNA detection technique in 1975 from his lab in England. It involved transferring DNA to a membrane and was dubbed a “Southern blot” in honor of its inventor’s name. Two years later, researchers at Stanford developed a similar RNA detection technique that was dubbed “Northern blot”. Two years after that, when Towbin transferred proteins to a membrane, it eventually became known as a “Western blot”.

What are the steps of a Western blot procedure?

1. Lysate/Cell Preparation. The number one thing you can do for success in your Western blot is appropriate sample prep. You will never be able to visualize your proteins if they remain trapped in your tissue or cells. You can read about our recommended lysis buffer here. You can purchase our lysis buffer here. Read about lysing your samples here. Read our lysate preparation protocol here.
2. SDS-PAGE. Separation of the proteins in your lysate by molecular weight is done through electrophoresis. Read all about the technique here.
3. Transfer. Getting the separated proteins out of the gel and bound to a membrane allows for easier detection. This is the procedure that I will highlight in this article.
4. Detection. This step involves incubating the transfer membrane in a solution containing an antibody to the protein of interest. When this antibody binds to the protein on the membrane, it can be detected with a chemiluminescent or fluorescent tagged secondary antibody allowing for visualization of the protein band.

Membrane Transfer Fundmentals

Why transfer the proteins to a membrane?

1. Ease of handling. Gels are fragile. We’ve all torn one. Membranes are hardier and are more easily manipulated.
2. Improved detection. Proteins are buried in the relatively thick gel. Getting them out of the gel and bound to the thin membrane allows for them to be more accessible to antibodies for detection.

Membranes? Tell me more.

Nitrocellulose and PVDF (polyvinylidene difluoride) are the membranes of choice for most Western blotting applications. Both membranes are microporous substrates that bind proteins to their surface through hydrophobic interactions. Here are the basic differences between the two:

Nitrocellulose. One of the first membranes used in Western blotting. It can bind protein at a capacity of 80–100 µg/cm².

• Pros:
  1. Lower background than PVDF (in part due to a lower binding capacity).
  2. Easier to block – less non-specific binding.
  3. Does not need to be pre-wet with methanol.
  4. Less expensive than PVDF.
  5. Better for low MW proteins whose binding is enhanced since methanol (which shrinks membrane pore size) is in the transfer buffer.
• Cons:
  1. Lower binding capacity than PVDF.
  2. Fragile – membrane can be easily chipped or cracked.
  3. Unsupported nitrocellulose can’t stand up to stripping and re-probing.
  4. Worse for high MW proteins – the methanol in the transfer buffer and the subsequent reduced pore size of the membrane can cause high MW proteins to precipitate.

PVDF (polyvinylidene difluoride). PVDF is a popular alternative to nitrocellulose due to its high binding and strength. Its binding capacity is 170-200 µg/cm².

• Pros:
  1. Resistant to solvents – can be easily stripped and re-probed.
  2. Higher binding capacity than nitrocellulose = higher sensitivity.
  3. Stronger and more resilient material – easier to work with.
• Cons:
  1. Higher background – (due in part to the higher binding efficiency).
  2. Needs to be activated with methanol.
  3. Usually more expensive than nitrocellulose.

Pore Size: Regardless of which membrane you use, you also need to consider pore size. Both types of membrane are microporous. The size of the pore determines the size of the protein that can bind without passing through. Membranes are available in different pore sizes, most commonly 0.2 um and 0.45 um. For most proteins, the 0.45 um size works well. For low MW proteins, <20 kDa, it is a good idea to use a membrane with a smaller pore size to keep your protein of interest from passing through the pores.

Transfer Mechanics

How do I transfer proteins from my gels to the membrane?

Most scientists use electroblotting to transfer their proteins from gels to membrane. A “transfer sandwich” of filter paper – gel – membrane – filter paper is placed between two electrodes. (In wet transfer systems, there is a sponge on either side of the sandwich.) The negatively charged proteins in the gel are pulled in an electric current toward the positively charged anode and into the membrane. Since the gel and membrane are sandwiched tightly during the procedure, the proteins maintain the separation they achieved during the SDS-PAGE electrophoresis. Prior to electroblotting, protein transfer could be performed through capillary transfer. This involves the same sandwich of filter paper, membrane, and gel, but relies on capillary action to pull transfer buffer from a lower reservoir, through the gel/membrane, and to the filter paper on the top, bringing the proteins with it and depositing them on the membrane. This method is not frequently used due to the lengthy procedure time, but it is decidedly less expensive than electroblotting because no fancy apparatus is needed.

Electroblotting Transfer Techniques: What’s the difference?

There are two primary electroblotting techniques: wet tank transfer or semi-dry transfer.

Wet-tank transfer – the “sandwich” described above – with sponges – is placed in between two electrodes and submerged vertically in a chamber containing transfer buffer. Some common tank transfer systems are shown here:

• Pros:
  1. Most widely used transfer system.
  2. More quantitative.
  3. Very high protein transfer efficiency – 80-100%.
• Cons:
  1. Lengthy transfer times – 1 hour to overnight.
  2. Lots of buffer, messy to assemble.
  3. Complex setup – lots of pieces, can introduce error.

Semi-dry transfer – the sandwich described above – without sponges – is placed horizontally in between two plate electrodes. The filter paper is wetted with transfer buffer, but there is no buffer reservoir or submerging of the transfer sandwich.

• Pros:
  1. Short transfer times – 10 minutes or less.
  2. Very little buffer, less waste.
  3. Simpler, less messy setup.
  4. High throughput, convenient.
  5. Sandwich components are sold preassembled.
• Cons:
  1. Patchiness and unevenness can be more of a problem due to the fast speed.
  2. Preassembled sandwiches are expensive.
  3. Lower protein transfer efficiency – 60-80%.
  4. Notorious for trapping bubbles, which cause spots on the membrane.

Essential Double-Check

How do I know if my protein has transferred?

Before starting your antibody incubation step, there are two staining procedures we recommend to make sure your transfer was successful.

1. Coomassie – incubate your gel in Coomassie blue and then destain to see if there is any protein left in the gel. It is likely that not all of the highest molecular weight proteins are out of the gel, but it will give you a good idea if the mid and low range proteins migrated out successfully.
2. Ponceau S – incubate your membrane in Ponceau S to visualize the banding pattern of the protein on your blot. This allows you to check for transfer efficiency as well as to highlight any problem areas on your gel.

Protein Visualization is Key to a Successful Western Blot

By Amy Archuleta

We advocate using Ponceau S stain to illustrate the efficiency of transfer from gel to membrane after removing it from the transfer apparatus.

Ponceau S staining is a wonderful way of illustrating the transfer efficiency as a whole, but also for highlighting any issues with small sections of the blot that could be problematic for that small portion, but that don’t negatively impact your entire blot.

These abnormalities (or normalities if your technique needs improvement) are invisible on a blot with no Ponceau S stain but become terrifyingly obvious after staining. They can happen in both semi-dry and tank (wet) transfer systems.
At right are some examples from a tank transfer set-up. The majority of the rat caudate lysate blot at right illustrates a beautiful transfer for mid-range MWs. There are two tiny bubbles on the far left, but these are easily avoided. A strange smudge near the bottom right, and some inconsistent spotting in the highest MWs illustrate that these sections of the blot are not ideal for bands that would show up in these specific areas.

Most Common Problem #1 – Bubbles

Bubbling can happen for many reasons. A bubble appears as a white spot on your membrane in the middle of the field of red Ponceau S stain. Bubbles are the result of air being trapped between the membrane and the gel during transfer.
Usually bubbling is minor, and if it is just a couple of bubbles like in the blot above, then I simply use a gel pen to circle them on the blot. Then when I use the blot later and the Ponceau S is washed away during blocking and incubation steps, I will still be able to see the compromised portion of the blot. This allows me to use the rest of the blot without worry.

Usual culprits:

1. Recently mixed transfer buffer. The agitation of mixing transfer buffer causes bubbling that can then get trapped.
   • Solution – Degas your transfer buffer. Let it sit for a while.
   • Solution – Mix fresh buffer without shaking – use a stir bar.
2. Pouring transfer buffer into transfer apparatus too quickly.
   • Solution – Pour the transfer buffer slowly into your tank to keep bubbling at a minimum. This is similar to the beer-against-the-side-of-the-glass technique.
3. Careless sandwich assembly. One of the most important things to do while setting up your transfer is to be observant and mindful as each layer is added. At each step in the process (filter paper – gel – membrane – filter paper) it is important to look for and remove any bubbles.
   • Solution: Rollers. Bubbles can be minimized by using a rolling device on the top filter paper after sandwich assembly to apply additional pressure and squeeze out bubbles. These rollers come in many different sizes and shapes, and you can even use a conical tube or cut serological pipette.
   • Solution: Pre-wetting transfer sandwich pads (sponges) with transfer buffer. If you are using a wet transfer box, adding dry pads on top of your submerged blot and filter paper can result in air sneaking into your sandwich. The pads act as a sponge, pulling buffer away from your blot.

Worst Case Scenario: Pictured below are blots of rat testes lysate run at the same time, in the same transfer apparatus, where the bubbling rendered the blots impossible to use. Actually, bubbling is only one of MANY issues on these blots! The left blot shows the bubbling as it appeared when emerging from Ponceau S stain. The right shows a similar blot on which I have circled all of the incriminating bubbles. Circling makes it very obvious that virtually no portion of this blot is usable.

Most Common Problems #2,3,4: Vertical Variation / Horizontal Waves / Smudged Banding

All three of these issues are very common. Most of them can be solved by tightly packing the transfer sandwich.

Vertical Variation:
This type of irregularity can appear differently (as seen in the image below) but is always indicative of non-uniform transfer of the protein in the gel to the membrane. The transfer sandwich must apply equal and firm pressure across the entire blot. Protein transfer efficiency will reflect any differences in pressure.

Horizontal Waves:
If the transfer sandwich has inconsistent pressure across the height of the membrane, horizontal waviness can appear. In the blot at right, it is clear that the gel was not tightly pressed against the membrane during transfer, allowing for buffer to pass between the two. At the top of the membrane, this resulted in the transferring protein “swimming” from one to the other and not maintaining its exact position. At the bottom of the membrane, what appears to be the edge of the gel is wavy and it is clear that swimming protein deposited itself beneath the gel’s edge.

Smudged banding:
Unsurprisingly, transfer sandwiches with insufficient pressure between the gel and the membrane can result in a loss of sharpness in the banding pattern. It is important to note that different lysates and different lysate preparations can also affect the banding pattern. However, it is always a good idea to try and minimize the smudge effect that can be caused by a too-loose transfer sandwich.

Usual Culprits:

1. 1. Insufficient tightness in the transfer sandwich. This is usually due to compressed sandwich pads. Over time and over many transfers, these pads become compressed just as any sponge would. Even after only a few uses, the compression is obvious.
      • • Solution: Replace your sandwich pads frequently.
        • Solution: Monitor the thickness of the pads and supplement your sandwich with additional filter paper (or an additional pad) if there is any pad compression.

       

1. Issues with the SDS-gel. No membrane is going show discreet band separation if the gel from which the proteins were transferred had a blurred resolution to begin with.
   • Solution: There are many things that can go wrong at this early step and are beyond the scope of this article. However, things to consider are: correct buffer concentrations and pHs, fresh components, and appropriate lysis buffer. If you are pouring your own gels, incomplete polymerization of your gel can also be to blame for poor resolution.

Common Problem #5 – Gel damage.

One of the main reasons that we transfer proteins from gels to membranes before probing with antibody is that gels are so fragile. When removing a gel from between the glass plates, placing it in the transfer apparatus, and assembling the sandwich, it is very easy to tear the gel. This doesn’t have to spoil your transfer, as the rest of the blot is probably fine. However once again it is vital to stain your membrane with Ponceau S to visualize and mark the problem area.

Usual Culprits:

1. Stuck gel. Gels are notorious for sticking to the glass plates when removing them from the SDS-PAGE apparatus.
   • Solution: Run a razor blade along the edges of your gel to separate it from the plate.
   • Solution: Use a squeeze bottle with water to shoot liquid in between the gel and the plate to further loosen the gel before trying to remove it from the plate.
2. Rough handling. Although they are relatively hardy, the truth remains that we’re dealing with thin, gelatinous sheets. Delicacy is the order of the day.
   • Solution: Be more gentle.

Common Problem #6 – Transfer Apparatus or Buffer Component Issues.

If you are sure that the above culprits aren’t the issue, check the function of your transfer apparatus and power source or your buffers.

Usual Culprits:

1. Apparatus Issues. There could be a problem with your set-up that is keeping you from success.
   • Solution: Black in back, red ahead! Make sure that your wires are attached to the correct electrode. If they are reversed, your proteins will be pushed away from the membrane and float away in the buffer.
   • Solution: Damaged Electrodes. Check the coating on your anode for wear, tear, and scratches.
   • Solution: Power Source. Is it still operational? Did you set it at the correct volts/amps?
2. Buffer Weirdness. Depending on your membrane and proteins of interest, your buffer may be to blame.
   • Solution: Reformulate your buffer. Research the buffer that works best in your system and with your membranes. Variables to address are methanol content, SDS content, salt concentration, and pH.
   • Solution: Remake your buffer. Mistakes were made. It happens. Sometimes just making fresh buffer can solve a world of problems.

To Sum it Up

There are a daunting number of variables that can affect a Western blot experiment. Tissue/cell lysis and preparation is a key component. Read about lysis buffers here, and goopy lysate here. SDS-PAGE efficiency is another hurdle. Read the basic science of SDS-PAGE here. After transfer to a membrane, antibodies can prove problematic for a variety of reasons. Read about antibody variability and validation here and here. Read about issues with secondary antibodies here. Read about the VERY basics of Western blotting here.

With all of these possible pitfalls, transfer efficiency is one issue that can usually be addressed quickly by three steps:

1. Taking time to prevent bubbles.
2. Packing your sandwich tightly.
3. Checking for transfer efficiency by staining with Ponceau S.

Happy Blotting!