I have six treatment groups (different cell types) with four biological replicates per group that I would like to compare protein levels in. I have come up with several experimental designs, each with pros and cons... For my first design, I thought to run four separate gels, each one containing 1 lane of each treatment group..For a second design, running six different gels, one for each cell type and having 1 lane per biological replicate. Trade-offs between the two being inter-gel variability and culture issues (different growing times, passage numbers, etc.)
Does anyone have any recommendations or thoughts on what the best way to go about this would be?
I think you should go with the second design since comparison of the each of the six cell types for the different conditions would be easier and analysis would be better.
Hi!! Did you try the incell western? You can use 96 wells in this assay!!!
If you have only 6 samples, as long as you use same percentage gel, you can run them on the same gel. You can leave lanes between cell types if you use 12 well combs or run couple dividing between the samples. If you want to probe with different primaries, you can still cut the membrane. It saves time and money. If you are happy running 6 gels, go ahead, there is nothing wrong in that, except time and money consuming.
Make sure you have included your internal ctrl group in each cell line experiments and then load them on the same gel, either according to the time of the cell lysate collection or all cell lysates to be compared together. Running the samples you want to compare with each other on the same gel is the best way of doing it unless you have an intra assay control that you run on every gel beside the rest of your samples. If you keep a good record of your total protein load and densitometry of your designated bands you can then normalize them first against your internal control & then against intra-assay control for comparing different gels. Just make sure your total protein load per well (ug/well) is the same among all your samples.
Good question!
first design!
in this way you could compare groups and consider gels as replicates.
Suggestion: load 1 samples in all gels to see how consistent your runs are
May be you can pool all 4 samples and use it as a control for every gel and then normalized the band with of each gel intensity to this band and check in all four gels
Running all samples on one gel might help you to quantify your protein levels more accurately. Also, your secondary antibody determines how accurate your quantification is. For example, AKP conjugated secondary might be better than HRP conjugated secondary.
I would go for the first design. The best would be to run all on one gel, but 24 samples might be too much. If you have each gel a replicate of the other, you can simply see if the trends are similar - as you want them to be. I do not know what protein will you use as a control for protein load, but you can normalize the gels (and the protein intensities) to each other using the intensity of the control protein.
May be can try with the second design ..just suggestion .or may be try both ..
this depends very much on the point you want to make with your experiment. Usually, you experiment is about the effects of the treatment, right? absolute protein levels will vary from cell line to cell line anyways.
Therefore, I would go for the second option and include a separate reference sample that is the same on each gel. Thereby, you can adjust the exposure time for the individual blots if absolute protein levels are interesting for you.
If you want to quantify a lot of samples (as mentioned above), it might be a good idea to set up an elisa for your protein (monclonal rat or mouse for capture and rabbit serum/polyclonal for detection works nicely in most cases).
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