I have two concerns:
1. In the ELISA that I'm using, I create a 4000 pg/mL high standard and 1:2 dilute down to a 62.5 pg/mL low standard. However, when I add the standards (as well as the samples) to my plate, they are added in 50 ul to another 50 ul of assay diluent (100 ul total volume per well). This makes me think that each sample/standard is AGAIN diluted 1:2. Is this dilution valid or do you ignore the 50 ul of assay diluent since each sample is treated that way?
2. As stated above, this ELISA assay begins with 50 ul of assay diluent in each well, to which I add 50 ul of sample/standards. My question is, what if you do not have 50 ul of sample? Is it valid to add amounts not equal to 50 ul? For example, if you only added 25 ul of sample, would you then just multiply by 3 at the end (75/25 = dil. factor of 3). Or do you not multiply by anything since you added your sample "neat"? And lastly, what if I add amounts less than 50 ul AND also use different amounts for different samples (e.g. 25 ul added for one sample, and 10 ul added for another sample). Is this okay so long as I take into account a relevant dilution factor? And what would that factor be?
Thanks a lot!
Cinnamon, for query 1 - you are correct, by adding your standards (in 50 ul) to 50 ul of diluent in the wells you are diluting down again by 1:2. The dilution is only valid if you want standards starting at 2000 pg/ml and going down to 31.25 pg/ml. However if you want that starting dilution, you should make your standards accordingly in the total 100 ul. Why are you putting your standards into another 50 ul of diluent?
For query 2, you can add less amounts of sample volume if you do not have enough, but you definitely have to take into account the dilution factor when analysing the results. Please see attached datasheet. Best practice would be to use the same dilution factor for each sample so perhaps stick with 10 ul/sample - which would give you a total volume of 65 ul - making this a 6.5 DF.
Hello Chelsea,
Thank you for your answer. I use the 50 ul of diluent simply because those are the instructions stated in the kit I'm using (maybe it equilibrates the well or something?). So based on what you said, I find it odd that they don't seem to take the extra dilution into account (see the kit instructions here: https://www.rndsystems.com/products/mouse-rat-leptin-quantikine-elisa-kit_mob00#assay-procedure (this is the simple version, the actual manual goes into more detail)). Maybe I'm missing something??
Re. your query 2 answer: This actually brings up another question I had. I, too, always had the impression that it would be best to use the same dilution for all samples, however in some of the work I'm doing, this is just not possible. For example, depending on the construct I'm transfecting cells with in vitro, I can get pretty wildly variable amounts of protein expression in the conditioned media afterwards (since it's secreted), which means I'm basically forced to used different dilution factors or half my plate will be out of the linear range. Do you think this is still okay? I suppose I could make plates of samples divided into those that need "high" or "low" dilutions, but then I would not be assaying the samples from the same experiment on the same plate.... What to do....?
Hi Cinnamon,
Re your query 2 response, Could you concentrate your conditioned media samples via concentration centrifuge tubes? Thus potentially bringing more of your samples into line - plus if you concentrate, you can use less sample to dilute them out to the linear range. I run multiple cancer cell lines and have to do this with certain cell line conditioned media, as protein expression is obviously different. Concentration has been successful and allowed me to run assays in the correct range.
Having said that, as long as you take into account your initial protein concentration and each dilution factor you've used, you have your answer.
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