Working with membrane proteins like WAKs in a Western blot can be frustrating, especially with their tendency to aggregate or denature poorly due to their hydrophobic regions. Here's how to make some adjustments that might solve your problem:

Avoid Boiling Your Samples

Boiling at 95°C can make membrane proteins like WAKs clump together. Instead, try heating them at 37°C for 10-15 minutes. This will gently denature the protein without causing it to aggregate, improving your chances of getting clean bands.

2. Switch to a Milder Detergent

While Triton-X is a good detergent, it may not always be enough for solubilizing tricky membrane proteins. Consider using digitonin or CHAPS—they're milder and can help protect the structure of your WAKs, allowing for better extraction.

3. Try a Membrane-Specific Lysis Buffer

Membrane proteins need different handling than cytosolic ones. A buffer like RIPA, which has both ionic and non-ionic detergents (e.g., SDS and NP-40), can improve solubilization. However, don’t go too heavy on the SDS, as it might interfere with antibody binding down the line.

4. Tweak Triton-X Concentration

Triton-X is good, but too much of it can lead to non-specific binding, contributing to the background noise you’re seeing. Adjust the concentration to somewhere between 0.1% and 1% to see what works best.

5. Reduce SDS in Your Sample Buffer

If you're using SDS in your sample buffer, reducing the concentration may help. Excess SDS can make membrane proteins aggregate, giving you smears or extra bands.

6. Boost Protease Inhibitors

Membrane proteins degrade faster, so upping your protease inhibitor concentration or adding phosphatase inhibitors (if you're worried about phosphorylation) might help stabilize your samples better.

7. Check Your Antibodies and Controls

Double-check that your HA and FLAG antibodies are specific and functional. You might want to run a dot blot first to make sure your antibodies are recognizing the tagged proteins properly. Also, ensure that your negative control truly has no expression of the target proteins to avoid misleading results.

8. Optimize Blocking and Washing Steps

Nonspecific bands often come from poor blocking or washing. Consider increasing your blocking agent concentration (e.g., 5% milk or BSA), and if background persists, extend the washes or use higher Tween-20 concentrations in your wash buffers.

9. Increase Protein Load

Membrane proteins don't always transfer efficiently to the membrane, which means you might need to load more protein on the gel to get a better signal. Make sure your transfer conditions are optimized too, particularly for large proteins.

10. Use a Gradient Gel

If you're dealing with proteins of various sizes, especially with YFP-fusions and WAKs, a gradient gel (4-20%) will help resolve them better, giving you clearer bands without smearing.

These tweaks should help improve your chances of getting the right bands. Experiment with a few of these adjustments and see how your results change. Keep me posted on how it goes

Faruq Sosanya Thank you so much i will try with suggestions you gave. Thank you