Western Blot

Lysis

Materials:

  • 10X Cell Lysis Buffer from CST

  • 200mM PMSF (200X concentrated)

  • Cell Scraper

  • Blunt 21 gauge needles

  • Lock-joint syringe

  • Cells to lyse

Note

  • If 10X cell lysis buffer will be continually used, it is recommended that the 10x buffer be kept at 4°C for 1-2 weeks. For longer periods of time, buffer should be stored at -20°C. Aliquoting of 10x buffer is recommended if many small experiments are to be performed.

  • PMSF is unstable in water and should be added to lysis buffers or other aqueous solutions just prior to use. Typically, a final concentration of 1 mM provides sufficient protease protection. Store lyophilized at RT for 24 months protected from direct light. Once in solution, PMSF can be stored at -20ºC for up to 3 months, protected from light.

Protocol:

Important

All reagents and lysates must be kept cold. In addition, keep plates on ice during lysis step.

Note

We usually use the 10X cell lysis buffer

  1. Cool microcentrifuge to 4°C.

  2. Dilute 10X Cell Lysis Buffer to a 1X solution using Elga water.

    • Can use either 10X lysis buffer or RIPA in -20°C “Western Reagents” box.

  3. Chill 1X buffer on ice and add PMSF just prior to use.

  4. Wash plate with ice-cold PBS to remove residual media.

  5. Add 400 uL of 1X lysis buffer to the 10cm dish of cells.

    Solution

    Vol for 1x10cm

    Vol for 3x10cm

    ELGA water

    358 µL

    1,170 µL

    10X cell lysis buffer

    40 µL

    130 µL

    200 mM PMSF

    2 µL

    6.5 µL

Note

BAL has also used 6-well plates to make cell lysate instead of 10cm dishes. Volume of 1x lysis buffer is scaled by surface area, so 68uL of lysis buffer is required per 6-well.

  1. Tilt to coat bottom of dish, then incubate on ice for 5 minutes.

  2. Scrape cells with a cell scraper and transfer lysed solution to a small centrifuge tube.

  3. Use 21 gauge needles to shear the cells 5-6 times (change out needle so 1 needle/sample).

  4. Spin extract 10-12 minutes at 14,000 x g (~11,500 rpm) in the 4°C microcentrifuge.

  5. Remove supernatant for use and aliquot.

  • You should have ~500-600 µL per 10cm dish (or about 70-80 uL per 6-well).

  • ~50-60 µL/aliquot is good because you can load 25 µL/well (12.5 µL lysate + 12.5 µL 2X Laemmli)

    so 50 µL will give enough for 4 wells (i.e. 4 antibodies).

  • BAL uses 20-25 uL/aliquot for lysates from a 6-well since 12.5 uL/well (6.25 uL lysate + 6.25 uL 2X Laemmli) has be sufficient for detecting proteins.

Note

  • Use prepared lysates as quickly as possible, and store for as short a time as possible.

  • Store lysates at -80°C for as long as possible. For lysates that will need to be kept around long term, transfer freshly prepared tubes to an available -80°C freezer to prevent degradation.

  • Lysates have a shorter shelf life when stored at -20℃; long-term storage at this temperature is not recommended. CST recommends that lysates are stored at -20℃ for no longer than 3 months.

  • Minimize your freeze/thaw cycles as much as possible. Instead, aliquot into smaller volumes.

Bradford Assay

Introduction

The Bradford Assay is used for protein concentration quantification. It utilizes Coomassie Brilliant Blue G-250 dye which binds to proteins and shifts the maximum absorption from 470nm to 595nm. This absorption at 595nm can be measured using the NanoDrop. The protein concentration can be estimated by creating a standard curve and using Beer’s law.

Materials

  • Bradford Assay Reagent

  • BSA Standard (2 mg/mL)

  • Lysis Buffer

  • Lysate Samples

Important

Ensure that the Bradford assay kit is compatible with the lysis buffer.

For example, the Bradford assay should be compatible with RIPA lysis buffer but is not compatible with the CST Cell Lysis Buffer due to high amounts of Triton.

See Bradford Assay Documentation for the full list of compatible reagents.

Protocol

  1. Remove Bradford Protein Assay Reagent from the refrigerator and gently mix by inverting the bottle several times.

  2. Aliquot the required volume of the dye reagent and allow it to equilibrate to room temperature before use.

  3. Prepare diluted BSA protein standards. Be sure to dilute the supplied BSA standard (2000 ug/mL) in the same buffer as the test samples (e.g. 1x lysis buffer). Use the tables below as a guide for preparing a set of protein standards.

For a working range of 100 to 1500 ug/mL:

Tube #

Buffer/Diluent Vol. (uL)

BSA Standard (2 mg/mL) Vol. (uL)

Final Protein Conc.

Blank

80

0

0

1

0

80

2000 ug/mL

2

20

60

1500 ug/mL

3

40

40

1000 ug/mL

4

50

30

750 ug/mL

5

60

20

500 ug/mL

6

70

10

250 ug/mL

7

75

5

125 ug/mL

8

79

1

25 ug/mL

For a working range of 1 to 25 ug/mL:

Tube #

Buffer/Diluent Vol. (uL)

BSA Standard (2 mg/mL) Vol. (uL)

Final Protein Conc.

Blank

800

0

0

1

790

10

25 ug/mL

2

792

8

20 ug/mL

3

794

6

15 ug/mL

4

796

4

10 ug/mL

5

798

2

5 ug/mL

6

799

1

2.5 ug/mL

7

799.5

0.5

1.25 ug/mL

  1. Combine each standard and unknown sample with the Bradford Reagent.

    • For a working range of 100-1500 ug/mL, pipette 1 uL of each standard or unknown sample into a labeled tube and add 20 uL of the Bradford Protein Assay Reagent and mix well.

    • For a working range of 1-25 ug/mL, pipette 10 uL of each standard or unknown sample into a labeled tube and add 10 uL of the Bradford Protein Assay Reagent and mix well.

  2. Incubate at room temperature for 10 minutes.

  3. On the NanoDrop, select the Proteins tab and then Bradford Assay.

  4. Enter the concentrations of each BSA standard and select the number of replicates.

  5. Measure the absorbance of of each BSA standard as directed by the NanoDrop to construct the standard curve.

  6. Measure the absorbance of each sample. The NanoDrop will automatically calculate the protein concentration for you based on the standard curve.

Protein Gel Casting

Modified for a Western Blot from this protocol.

Required stock solutions

  • 40% Acrylamide stock solution: Solution of monomers for gel polymerization.

    We find it cheaper to buy premixed 40% stock solution, with a acrylamide:bis-acrylamide ratio of 29:1 (3.3%). Stocks with a 37.5:1 ratio also work, and are typically used for resolving larger proteins.

  • 3x bis-Tris gel buffer: Ion buffer used in gel casting.

    Component

    Concentration

    g/L to final concentration

    bis-Tris

    1 M

    209.242

    HCl

    Add to pH 6.5-6.8

  • 10% APS: One of the polymerization initiators. Only a small quantity needs to be prepared; each gel only requires 35 uL. Make fresh each time by dissolving in water.

    Component

    Concentration

    g/L to final concentration

    Ammonium persulfate

    10%

    100 (For example: 10mg/100uL or 100mg/1mL)

Casting protocol

Warning

The acrylamide monomers used here are toxic. Read the SDS.

Perform polymerization steps with a lab coat in a fume hood, and collect rinse waste in a waste container.

  1. Prepare 1X resolving and stacking buffers. These buffers can be stored in the refrigerator for several weeks. Recipes given here for enough for 2 0.75mm gels.

    Resolving buffer: ~3 mL per gel (6.5 mL total). Final acrylamide concentration depends on desired protein size:

    Protein Size

    Gel %

    Vol 40% Acrylamide Stock

    Vol DI Water

    Vol 3x bis-Tris gel buffer

    4-40 kDa

    20%

    3.25 mL

    1.05 mL

    2.2 mL

    12-45 kDa

    15%

    2.44 mL

    1.86 mL

    2.2 mL

    10-70 kDa

    12.5%

    2.03 mL

    2.27 mL

    2.2 mL

    15-100 kDa

    10%

    1.63 mL

    2.67 mL

    2.2 mL

    25-100 kDa

    8%

    1.30 mL

    3.00 mL

    2.2 mL

    Stacking buffer: ~1.2 mL per gel (2.5 mL total):

    Component

    Volume

    Final concentration

    3x bis-Tris gel buffer

    0.83 mL

    1x

    40% Acrylamide stock

    0.32 mL

    5%

    DI water

    1.36 mL

    Bromophenol blue

    50 uL (enough to give color which helps when loading)

Gel casting setup

In-lab, we have the ability to cast two gels simultaneous; this is recommended even if you only need one, so that you have a backup in case of pouring mishaps. Our gel runner also requires two poured gels to properly seal.

  1. Locate two 0.75mm spacer plates and two short glass plates.

  2. Use ethanol and a Kimwipe to clean both glass surfaces.

  3. Assemble them in the green alignment device.

  4. Lock the two gels into the transparent gel pouring device.

Resolving gel

Tip

The resolving gel can polymerize within a just minute or two, especially at higher percentages of acrylamide. Therefore, pour the gel quickly using a P1000 pipette.

It is best to pour the gel from the edges of the gel mold to avoid bubbles.

Get ~10 mL isopropyl alcohol (IPA) ready before pouring the resolving gel to help keep the gel interface straight and level.

  1. Prepare fresh 10% APS. 1 gel requires ~35 uL so if making 2 gels, prepare ~100 µL (10 mg).

  2. Measure 6.5 mL of 1X resolving buffer per gel to pour.

  3. Add 50 uL of 10% APS per gel, mix well.

  4. Add 20 uL TEMED, mixing quickly (don’t pipette mix, just flip it x3 manually to mix).

  5. Pour both gels to the resolving gel height (3 mL per gel, 1,000 µL at a time).

  6. Ideally there shouldn’t be bubbles, but if so, lightly tap and tilt the gel to remove

  7. Once done pouring, quickly but carefully fill the remaining height with IPA, making sure the gel-water interface stays undisturbed. This is to ensure the resolving-stacking interface is straight and level.

  8. Wait for the polymerization reaction to finish (noticeable by a change in refractive index).

  9. Drain the IPA by tilting the gel past 90 degrees, and wicking away with a Kimwipe.

Stacking gel

  1. Measure 2.5 mL of 1X stacking buffer to pour.

  2. Add 20 uL of 10% APS, mix well.

  3. Add 10 uL TEMED, mixing quickly (don’t pipette mix, just flip it x3 manually to mix).

  4. Add 1,000 µL of stacking gel into each gel.

  5. Insert the comb into the top very carefully, one edge at a time to avoid bubbles. The stacking gel will overflow.

  6. If any bubbles, pop comb slightly up near problem area and use remaining buffer to fill before closing again.

  7. Wait for the stacking gel to polymerize.

  8. Rinse with water or IPA (evaporates faster) to remove unpolymerized acrylamide.

  9. If removing the combs prior to storage, slowly remove the comb, ensuring that wells are not broken.

Loading and Running the Gel

Modified for a Western Blot from this protocol.

Solutions required

  • 20x MES-SDS running buffer stock solution: Suitable for separating proteins with a molecular weight less than 75 kDa.

    It is also generally cheaper to order this as a pre-mixed 20x stock solution. If you need to make it yourself, the recipe is:

    Component

    Final concentration

    g/L to final concentration

    MES

    1 M

    195.2 g

    Tris

    1 M

    121.13 g

    EDTA

    20 mM

    5.845 g

    SDS

    2%

    N/A

  • 200x running buffer reductant: Ensures that the gel remains under reducing conditions when run. Add directly to 1x running buffer before filling the gel tank. Dissolve sodium bisulfite in DI water.

    Note

    Dilute sodium bisulfite solution loses effectiveness in ~2 days so spike in fresh each time.

    This helps because although β-mercaptoethanol in the Laemmli buffer is a strong reductant that prevents crosslinking via reduction of disulfide bonds, over time it can degrade.

    Component

    Final concentration

    g/L to final concentration

    Sodium bisulfite

    1 M

    104.061 g

  • 200 mM Tris-HCl stock: Dissolve components in DI water.

    Component

    Concentration

    g/L to final concentration

    Tris-HCl

    200 mM

    31.52 g

    NaOH

    Add to pH 6.8

  • 20% SDS stock: At low temperatures, the SDS may fall out of solution. Therefore, warm in a water bath to dissolve. Mix well before transferring.

    Component

    Concentration

    To make final concentration

    Sodium dodecyl sulphate

    20%

    2g / 10 mL DI water

  • 0.1% bromophenol blue: 1 mg / mL

  • 2x Loading Buffer (Laemmli Buffer): Used to denature and solubilize protein samples. Can be stored at 4°C.

    Component

    Final concentration

    Volume

    200 mM Tris-HCl stock

    100 mM

    5 mL

    Glycerol

    20%

    2 mL

    20% SDS stock

    4%

    2 mL

    0.1% bromophenol blue stock

    0.01%

    1 mL

    2-mercaptoethanol

    10%

    1.1 mL

Warning

2-mercaptoethanol smells awful; always add it inside a fume hood.

2-mercaptoethanol is hazardous waste and should be disposed of in the proper waste container.

Running procedure

  1. Add 2x Laemmli Buffer to an equal volume of lysate in PCR tubes. 50-60 µL is good for ~4 lane (need 12.5 µL lysate/lane) This is recommended unless the online antibody datasheet indicates that non-reducing and non-denaturing conditions should be used.

  2. Use a PCR machine to reduce and denature the lysate samples at 95℃ for 5 minutes (use 4℃ hold at end to keep cold).

  3. Dilute enough 20x MES-SDS running buffer to fill the gel tank, adding fresh 200x running buffer reductant if a gel has not been recently run.

  4. Place a prepared bis-Tris protein gel in the gel-runner. Fill both chambers with the prepared 1% MES-SDS running buffer. Fill the inner chamber to the top of the stacking gel, and the outside chamber to the top of the resolving gel. You will need about 1 liter of the 1% MES-SDS running buffer.

  5. Carefully load equal amounts of protein samples, including 5 µL of a protein ladder, into the wells of the gel. Each well can be loaded with a maximum of 25 uL. 20-30 ug of total protein from cell lysate is generally used unless further optimization is needed for the desired protein(s).

    • The protein ladder is in the -20℃ fridge in the restriction enzyme ice box

    Note

    BAL has found loading 12 uL of denatured lysate in Laemmli buffer per well is sufficient to detect most proteins.

    BAL tried freezing Laemmli buffer-denatured lysate at -20℃ and it worked for Western

    Tip

    Choose an asymmetric loading pattern so if the gel is flipped over, you will still know the order of your samples.

    Warning

    The glass gel holders have directionality! If your gel isn’t reaching 30 mA check that the open side is facing inwards.

  6. Run the gels at constant current, about 30 mA (~43V) per mini-gel for approximately 125 minutes. The dye band runs around 3-5 kDa, so it is typically ok to run the dye band to the bottom of the gel unless very small proteins are of interest.

    • Rinse gel holder and runner with water to help reduce smell

    Note

    BAL has run the gel for up to 140 minutes and found this helps separate out some of the larger proteins such as pERK which has bands at both 42 and 44 kDa.

  7. Pour DI water into a plastic tray (tip box lid), about half a centimeter deep.

  8. Very carefully separate the gel plates without breaking the gel. The gel will stick to one side or the other.

  9. With a razor blade, cut off the stacking portion of the gel while still on glass.

  10. Invert the plate/gel over the water and “convince” the gel to fall into the dish. It can help to put the gel and plate into the water and let the solution help the gel release. Using a green gel scraper can also help with this process.

  11. Place the gel on a rocker for 2-5 minutes to remove excess free proteins.

Coomassie Staining

Solutions required

  • Coomassie staining dye: When preparing this dye, pour the 10% methanol first, using it to dissolve the R-250. Then, add water. Add the glacial acetic acid last to prevent aggregation.

    Component

    Final concentration

    Amount per 1 liter

    Coomassie R-250

    0.2% (2g/L)

    2g

    Methanol

    10%

    100 mL

    Water

    80%

    800 mL

    Acetic acid

    10%

    100 mL

  • 10% Acetic Acid: Used as a destain solution.

    Warning

    Do not microwave pure acetic acid.

Procedure

  1. Drain the water from the dish without dropping your gel in the sink, and cover with ~0.5 cm of Coomassie staining dye.

  2. Place the gel in stain in the microwave and microwave on high until the solution just begins to boil (this step greatly accelerates the procedure and allows you to see you bands in a minute or so). This only takes 20-30 seconds in the microwave.

  3. Remove from the microwave and place on a rocker for a few minutes. Once you see the gel filled with Coomassie, it’s done.

  4. Drain the Coomassie and cover the gel with water, rock for about 5 minutes, drain.

  5. Cover with 10% acetic acid, place a couple folded Kim-wipes over the gel, and microwave again until the solution begins to boil (20-30 seconds).

  6. Remove from microwave and rock to remove Coomassie not bound to protein. If there is excess stain, replace the 10% acetic acid and Kim-wipes and continue to rock until the gel is clear with dark purple protein bands.

Transferring the protein from the gel to the membrane

Tip

Proteins in the gel can be transferred to a membrane using the iBlot2. It is recommended to watch this iBlot2 tutorial video to learn how to use this device. The iBlot2 manual is another helpful resource for learning to use the iBlot2 and contains more detailed instructions.

  1. Open the lid of the iBlot2 device using the latch. Ensure the blotting surface is clean. Wipe down electrical contacts.

  2. Unseal the iBlot™ 2 Transfer Stack.

  3. Separate the Top Stack and set it to one side of the bench with the transfer gel layer facing up.

    Keep the Bottom Stack in the transparent plastic tray. Top and bottom stacks are divided by a separator. Ensure the membrane is not stuck to the separator and is with the bottom stack.

  4. Place the Bottom Stack with the plastic tray directly on the blotting surface.

  5. Ensure there are no bubbles between the membrane and the transfer stack. Remove any trapped air bubbles using a roller such as a plastic conical.

  6. Carefully separate the glass plates surrounding the gel so the gel is left on one of glass plates. Using a razor blade, cut off the stacking region of the gel. Then carefully remove the gel from the glass into a dish filled with 1 cm of DI water.

  7. Place the pre-run gels on the transfer membrane of the Bottom stack. Note: two gels can fit onto a single membrane.

  8. Use a roller to remove any air bubbles between the gel and the membrane.

  9. Soak the iBlot Filter Paper from the transfer stack in a clean container of deionized water.

  10. Place the presoaked iBlot Filter Paper on the pre-run gel. Use a roller to remove any air bubbles between the filter paper and gel.

  11. Remove and discard the white plastic separator from the Top stack.

  12. Take the Top Stack from the bench and place it on top of the presoaked filter paper with the copper electrode facing up (and transfer gel layer facing down). Remove any air-bubbles using a roller.

  13. Place the iBlot™ 2 Absorbent Pad on top of the iBlot™ 2 Transfer Stack such that the electrical contacts are aligned with the corresponding electrical contacts on the blotting surface of the iBlot™ 2 Gel Transfer Device.

  14. Use the Blotting Roller to flatten any protrusions in the transfer stack.

  15. After assembling the iBlot™ 2 Gel Transfer Stack, perform blotting within 10-15 minutes of assembling the stacks with the gel.

  16. Gently close the iBlot™ 2 Gel Transfer Device lid by pressing down with two hands on the sides of the lid. Make sure the latch is secure. Do not forcibly push the lid when closing, because it can cause the transfer stack or metal contacts to shift out of position.

    Note

    If the iBlot2 device says the transfer stack is not detected, try opening and closing the lid until you are able to start the program. Cleaning the electrical contacts before and after each use may help with this issue.

  17. Ensure that the correct Method is selected.

Method

Voltage

Default Run Time

Run Time Limit

P0
20 V for 1 min
23 V for 4 min
25 V for 2 min

7 min

13 min

P1

25 V

6 min

10 min

P2

23 V

6 min

11 min

P3

20 V

7 min

13 min

P4

15 V

7 min

16 min

P5

10 V

7 min

25 min

Note

See page 17 of the iBlot2 manual for more detailed information about running parameters.

Transfer protocol P0 with default run time has worked previously when blotting for Beta-actin, WT-p53, and RAS.

For proteins from 30 to 150 kDa method P0 for a 7 minute run time is recommended. For proteins >150 kDa methods P0 or P3 with a run time of 8-10 min is recommended.

  1. Touch the Start icon on the screen to begin the transfer.

  2. At the end of the transfer, the current automatically shuts off and the iBlot™ 2 Gel Transfer Device signals the end of transfer with repeated beeping sounds, and a message on the digital display.

  3. Touch the Done icon to stop the beeping.

  4. To obtain good transfer and detection results, open the device and disassemble the stack within 30 minutes of ending the blotting procedure.

  5. Open the lid of the iBlot™ 2 Gel Transfer Device.

  6. Discard the iBlot™ 2 Absorbent Pad and Top Stack.

  7. Carefully remove and discard the gel and filter paper. Remove the transfer membrane from the stack.

  8. If needed, cut the membrane with a razor blade or scissors to separate the regions corresponding to each gel.

Antibody Staining

Solutions required

  • 10X Tris-Buffered Saline (TBS): Add ~450 mL of DI water to dissolve the Tris and NaCl. Adjust to a pH of 7.6. Then add the remaining DI water to reach a final volume of 500 mL.

    Note

    Took ~8-9 mL 12N HCl to get to pH ~ 7.6

    Component

    Final concentration

    Amount Needed

    Tris-base

    200 mM

    12 g

    NaCl

    1500 mM

    44 g

    DI Water

    To 500 mL

  • 10% Tween20:

    Component

    Final concentration

    Amount Needed

    Tween20

    10%

    1 mL

    DI Water

    9 mL

    Note

    Larger volumes of Tween20 are easier to measure because it is very viscous.

  • 1x Tris-Buffered Saline / Tween (TBST):

    Component

    Final concentration

    Amount Needed (50 mL)

    Amount Needed (1 L)

    10X TBS

    1X

    5 mL

    100 mL

    10% Tween20

    0.1%

    0.5 mL

    10 mL (or 1 mL Tween-20)

    DI Water

    To 50 mL

    890 mL (or 900 mL)

  • Blocking Buffer:

    Component

    Final concentration

    Amount Needed

    Milk Powder

    5%

    2.5 g

    10% Tween20

    0.1%

    0.5 mL

    10x TBS

    1X

    5 mL

    DI Water

    To 50 mL

  • 10% Blocking Buffer: For diluting primary and secondary antibodies.

    Component

    Final concentration

    Amount Needed

    Blocking Buffer

    10%

    5 mL

    1x TBST (TBS/0.1% Tween-20)

    45 mL

Staining Procedure

Note

A 10 cm dish works well for the wash steps.

  1. Wash the membrane with DI water for 5 minutes using agitation.

  2. Block the membrane with blocking solution for 30-60 minutes at room temperature with agitation. Alternatively, block overnight at 2-8°C. (NW does 60 min at RT).

  3. Incubate the membrane with 4 mL/10 cm of primary antibody diluted (at manufacturer’s recommended dilution) in 10% blocking solution overnight at 2-8°C.

  4. Wash the membrane 3 times for 10 minutes each in TBST using agitation to remove any unbound primary antibody.

  5. Incubate blot with 4 mL/10 cm of secondary antibody HRP-conjugate at a 1:10,000 dilution (or at the manufacturer’s recommended dilution) for 30 minutes to 1 hour at room temperature using agitation. (NW does 1 hr at RT)

    • Can even go down to 1:50,000 for 2nd ab HRP-conjugate

  6. Wash the membrane 6 times for 5 minutes each in TBST to remove any unbound secondary antibody conjugate. It is crucial to thoroughly wash the membrane after incubation with the HRP enzyme conjugate.

  7. Prepare the SuperSignal West Femto Substrate working solution by mixing equal parts of the Substrate and Stable Peroxide components (e.g. 5 mL substrate with 5 mL stable peroxide). Use a sufficient volume (~3 mL/10 cm) to ensure the blot is completely wetted with the substrate and does not become dry.

    Note

    The working solution is stable for up to 6-8 hours at room temperature.

  8. Incubate the membrane with the substrate working solution for 5 minutes.

  9. Remove the blot from the working solution and place it in a labeled, clear plastic bag, and remove excess liquid with an absorbent tissue.

  10. Image the blot using chemiluminescence. The membrane does not need to be removed from the clear plastic bag for imaging. The Niles Lab in BE has a ChemiDoc Imaging System that they let us use, and images can be transferred using a USB flash drive.

  11. Blot quantification can be done using the Gel Analyzer tool in ImageJ.

Note

Use colorimetric for a black/white photo that you can merge with the chemiluminescence photo

Note

NW uses optimal auto-rapid as default

Note

BAL has used a fluorescent secondary antibody instead of chemiluminescence since the ChemiDoc Imaging System can detect various fluorescent channels. This method seems to be slightly less sensitive and requires a high concentration of antibody for protein detection. Anti-rabbit secondary antibodies seem to bind to the protein ladder. Keep blots protected from light after adding the secondary antibody during staining. Online protocols suggest that it is important to make sure the membrane is dry before imaging.

BAL has been able to re-probe a fluorescent blot for chemiluminescence. You can rewet the membrane by first using 70% ethanol and then washing with TBST because the PVDF membrane does not absorb aqueous solutions uniformly unless pre-wet. Once rewet, you can proceed to re-probe the membrane.