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Troubleshooting for Quantitative Western Blots

The best transfer conditions, membrane, and blocker for experiments depend on your antigens and antibodies. For data examples and troubleshooting tips, get the Good Westerns Gone Bad technical note.

Uniformly distributed high background

Membrane contamination

Always handle membranes carefully and with clean forceps. Before you use them, clean dishes, bags, or trays for incubations with methanol.

Blocking conditions

Different primary antibodies will react differently in different blocking buffers. Use the Blocking Buffer Optimization kit to determine the best blocking buffer for your primary antibody.

Cross-reactivity of antibody with blocker, especially milk

Don’t use milk for blocking, as milk typically contains IgGs that cross-react with anti-goat secondary antibodies. Instead, try the Odyssey® Blocking Buffer.

Lack of detergent

Add Tween® 20 to primary, secondary, and wash steps at 0.1 – 0.2%.

For PVDF membranes, add SDS to secondary antibody incubation (not primary antibody incubation or wash step) at 0.01 – 0.02%.

For nitrocellulose membranes, do not add detergent to the blocking step.

With IRDye® 680LT secondary antibodies, use SDS (0.01 – 0.02% final concentration) and Tween 20 (0.1 – 0.2% final concentration) during the detection incubation step.

Not enough washing

Wash 4 times, for 5 minutes each wash. If background persists, increase the number of washes and buffer volume. Make sure that 0.1% Tween 20 is present in buffer. Excess Tween 20 (0.5 – 1%) may decrease signal.

Antibody concentrations too high

Do not add secondary antibodies at concentrations higher than 1:10,000 dilutions, as high background will result. A good starting dilution is 1:20,000. Use the MPX™ blotting system to optimize primary and secondary antibody dilutions. Get the One-Blot Western Optimization: Using the MPX Blotting System technical note for more information. For IRDye secondary antibodies, dilute between 1:10,000 – 1:40,000.

Old secondary antibody

IRDye secondary antibodies are stable for 3 months at 4° C. After 3 months, the effectiveness of secondary antibodies will decrease, and background may increase.

Not enough antibody

Increase antibody volume so entire membrane surface is sufficiently covered with liquid at all times. Use enough reagents to prevent areas of the membrane from drying out during incubations and washes. Gently agitate during every antibody incubation.

Background from PVDF membrane

Use low-fluorescence PVDF membrane, and confirm uniform membrane background before you transfer. LI-COR prescreens Immobilon-FL PVDF membrane kits for quality control.

Blocking with BSA

Blocking solutions containing BSA may cause high membrane background and nonspecific antibody binding for near-infrared Western blots. Avoid BSA for blocking. For the best sensitivity, try the Blocking Buffer Optimization Kit to compare different blocking conditions.

Uneven, blotchy, or speckled background

Blocking multiple membranes together in small volume

Your data will not be quantitative if you incubate multiple membranes together in one container. If you must block multiple membranes in the same dish, use enough blocking buffer for all membranes to move freely and fully contact the blocker.

Partially dry membrane or membrane not fully wet

If using PVDF, pre-wet the membrane in 100% methanol until it becomes translucent gray instead of opaque white. Wet membrane in PBS or TBS for 5 minutes or until uniform in color. Then place the membrane in blocking. Pre-wetting is necessary to help the PBS or TBS interact with the membrane, because PVDF membranes are very hydrophobic.

If using nitrocellulose, wet membrane in PBS or TBS for 5 minutes, or until uniform in color, before blocking.

After blocking, keep membrane completely wet at all times during the blotting process. This is especially important if you’re planning to strip and reprobe the blot.

Membrane handling

Avoid touching membranes with gloved or ungloved hands, as fingerprints will fluoresce on the Odyssey imaging system. Always handle membranes with clean forceps, free of any contaminants or antibody solutions. Clean forceps with methanol before using or after dipping them into an antibody solution (especially dye-labeled secondary antibody). Dirty forceps deposit dye on the membrane that you can’t wash away.

Contaminated dishes, boxes, or trays

Clean dishes, boxes, or trays with methanol before using them for incubation.

Dirty scanning surface or silicone mat

Clean scanning surface and mat carefully before each use. Dust, lint, and residue will cause speckles.

Marks on membrane from incompatible marker or pen

Use pencil to mark membranes. For nitrocellulose membranes, you can also use an Odyssey® pen .

Dirty transfer pads or transfer box

Transfer pads in wet tank systems and transfer boxes accumulate residue after frequent use that can cause speckles on Western blot membranes. Clean transfer pads and transfer boxes by soaking them in 100% methanol for 10 minutes.

Dirty imaging system

Always clean your imaging system before you image. For the Odyssey CLx, Odyssey Sa, and Odyssey Classic imaging systems, clean the glass surface with an alcohol-based cleaner before imaging.

For the Odyssey Fc imaging system, avoid using imaging trays that have been used for Coomassie stained gel imaging. Clean trays with an alcohol-based cleaner before you image.

Weak or no signal

Not enough antibody used

Primary antibody may have low affinity for your target. Increase amount of antibody or try a different supplier.

Extend primary antibody incubation time. Try 4-8 hours at room temperature, or overnight at 4 °C. Do not reuse antibody.

Increase amount of primary or secondary antibody, optimizing for best performance. Try the MPX Blotting system to optimize antibody concentration. For details, get the One-Blot Western Optimization: Using the MPX Blotting System technical note.

Degraded antibody

Primary and secondary antibodies can lose reactivity from improper or extended storage. Examine the product’s shelf life, and consider replacing with fresh antibody stocks. Avoid reusing antibody.

Signal washed away by too much detergent

Decrease Tween 20 in diluted antibodies. Recommended Tween 20 concentration is 0.1 – 0.2%.

For PVDF membranes, recommended SDS concentration is 0.01 – 0.02% during the secondary antibody incubation step (in addition to Tween 20). Some antibodies require an even lower concentration.

For nitrocellulose membranes, do not add SDS to any steps.

Unsuitable blocking buffer for experiment

Your primary antibody may work much better with a different blocking buffer. For tips on how to choose an appropriate blocker, get the Odyssey Western Blot Blocker Optimization for Near-Infrared (NIR) Detection protocol.

Inappropriate antigen amount loaded

Loading too little or loading too much protein sample will decrease antibody sensitivity. Loading too little sample results in not enough antigen present. Loading too much sample can cause your target antigen being masked by other proteins or antibody hindrance. Determine the optimal loading concentration by performing a serial dilution of your target of interest.

Try loading a dilution series that ends with the original amount of antigen that didn’t produce enough signal. This helps you determine the best signal with the lowest amount of antigen. You can also use the narrowest possible well size to concentrate antigen.

Protein did not transfer well

Check transfer buffer choice and blotting procedure. For more details, get the Protein Electrotransfer Methods technical note.

Wet tank transfer is the gold standard for protein transfer. Validate your transfer method to ensure your target antigen transfers under the preset conditions. The amount of methanol in the transfer buffer, timing of gel presoak, choice of membrane, voltage, and length of transfer can all change how much protein transfers to the membrane.

A pre-stained molecular weight marker can help you monitor transfer. You can also use REVERT™ Total Protein Stain to stain membranes post-transfer to monitor transfer efficiency. Stain gels with REVERT or Coomassie blue after transfer to see if the gel retained any proteins.

Protein lost from membrane during detection

After transfer is complete, dry your membrane before you block. Drying the membrane allows proteins to bind tightly to the membrane, preventing potential signal loss.

For PVDF membranes, re-activate membranes with methanol and rinse with water before blocking.

Proteins not retained on membrane during transfer

SDS in transfer buffer may interfere with binding of transferred proteins, especially for low molecular weight proteins. Try reducing or eliminating SDS. Note: presence of up to 0.05% SDS does improve transfer efficiency of some proteins.

Air-dry membranes completely for 1 hour (or 10 minutes at 37 °C) after transfer, to make binding irreversible.

Transfer of large proteins (>140 kDa) often requires lower methanol concentrations (10%) and possibly the addition of SDS to 0.05%. To get complete transfer of larger proteins, extend transfer times.

Small proteins may pass through the membrane during transfer (“blow-through”). To get good membrane retention of these smaller proteins, use higher methanol concentrations (30 – 40%) and lower voltage transfer.

Over-blocking the membrane

Extended blocking times can mask antigen and decrease signal intensities. Avoid blocking membranes for more than 1 hour. Block membranes at room temperature. For the best experimental reproducibility, block membranes at consistent times and temperatures.

Non-specific or unexpected bands

Unsuitable blocking buffer for experiment

The blocker you use may affect background bands. Use the Blocking Buffer Optimization Kit to try a different blocker.

Antibody cross-reactivity in a two-color Western blot

In two-color Western blots, antibody cross-reactivity is always a possibility. To avoid cross-reactivity:

  • Detect the two secondary antibodies on two separate blots first. See what each channel looks like individually before you detect the two secondary antibodies together on the same blot, to help you know what bands to expect and where to expect them.
  • Use secondary antibodies from the same host species to avoid potential cross-reactivity.
  • Avoid using mouse and rat primary antibodies together if possible. Because the species are so closely related, anti-mouse will react with rat IgG to some extent, and anti-rat with mouse IgG. Sheep and goat antibodies may also cross-react.
  • Use only highly cross-adsorbed secondary antibodies, like the IRDye® secondary antibodies.
  • Use less secondary antibody to minimize cross-reactivity – a good starting dilution is 1:20,000. Dilute between 1:10,000 – 1:40,000 for optimal performance.

Too much antibody

  • Use the supplier’s lowest recommended amount of antibody.
  • Reduce antibody incubation time to 1 hour at room temperature.
  • Dilute antibodies in the same blocking solution that you used to block the blot. IRDye® secondary antibodies dilutions can be increased up to 100,000.
  • Include Tween 20 at 0.2%.
  • For PVDF membranes, add or increase SDS in diluted secondary antibodies.

Signal bleed from one channel to the other

Check your fluorescent dye. Fluorophores like Alexa Fluor® 750 may appear in both channels (700 nm and 800 nm), and are not recommended for use with Odyssey® imaging systems.

If signal in one channel is very strong (near or at saturation), it may cause signal in one channel to bleed to the other channel. Minimize this by using a lower scan intensity setting in the channel that had strong signal. If you have an Odyssey CLx imaging system, try the AutoScan mode.

Reduce signal by reducing the amount of protein loaded or the amount of antibody. A dilution series of each will also help you save on sample and antibody.

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