high slope (b-value) of standard curve - (Jul/05/2013 )
I am running an elisa using Bethyl IgG-Fc capture antibody @ 10ug/ml coating (CBC coating buffer per Bethyl), using a medium binding plate. The standard is a Bethyl IgG human reference serum. I am starting the dilution of the std at 500ng per the protocol & diluting 2-fold. The secondary IgG aby dilution is 1:15,000 (all 3 components come as a kit). I am coating the rest of the plate with a specific protein for which I am trying to quantify antibody amounts and running random serums in them.
The OD of the IgG STD start at about 3.0, but once they come to the well that the conc. is at 125ng (3rd well), the ODs drop way past 2-fold & continue to drop fast resulting in a high slope of the standard curve & very little "plottable" points. The ODs for the wells coated with the specific protein (random serums run) dilute out just fine, it's only the STD column that is giving the issue, so I can't quantitate until this is resolved.
I have tried:
-starting the IgG STD at 250ng & 1000ng, the same thing happens once the dilution of it hits 125ng, no matter the starting conc.
-today I am trying a high binding plate with various 2nd ABY conc.
-today I am trying the same medium bind plates and increasing the 2nd ABY diln from 15,000 to 5,000
Why is this only happening in the capture ABY/IgG STD column & not the specific protein coated wells? I am using the same conc. of 2nd aby across the plate.
I am not getting background in my blank wells.
Just read the results using the high-binding plates. The ODs are much better, not getting the huge decrease is ODs, but slopes are still higher than desired (1.5ish). Is this just a balance of an antigen/aby amount available to bind ratio? or maybe too much or not enough incubation times, but at which step?
thanks for any insight
I am having difficulty in understanding what you are trying to measure and what the format of the assay is.
In order to compare quantities of unknown against your human IgG standard curve, the entire plate should be coated with the same thing (i.e bethyl anti-IgG Fc or your specific protein, but not both). It seems that you are running two different assays on one plate:
assay 1 (curve): capture with anti-human IgG Fc, human IgG standard, detect with another anti-human IgG-HRP (or other conjugate)
assay 2 (samples): capture with specific protein (presumably some viral or vaccine antigen), samples containing unknown titres/quantities of anti-antigen antibodies, detect the antibodies with the anti-human IgG-HRP
As the two formats rely on different coating efficiencies and binding kinetics, it is not surprising that the two curves exhibit lack of parallelism (can not be super-imposed).
Unless I am missing something, the entire plate should be coated with your specific protein, and you need a reference serum (or purified standard) containing or consisting of human anti-antigen IgG to use as your standard.
Let us know how you get on, or if you need any more help with this assay
good luck!
Yes, we are coating with 2 different proteins, an IgG capture aby for the STD curve column (a human IgG STD serum w/ known amount of IgG) & specific protein in the other columns- we are only looking for a relative IgG concentration in compared to the amount of expected IgG in the STD serum, not necessarily specific.
Anyway, with the high bind plates, I can get the STD curve to come down half-way decent for the IgG STD with slopes about 1.5 (no range errors given in SMPro, but higher b values than i'm used to seeing), but the serum samples run in the wells coated with specific protein have really high CVs. We are diluting 2-fold & the starting ODs are about 2.5ish, but fall slowly, making the adjusted results in SMPro have a huge range of titers & creating high CVs (40-60%).
I am using a block step before adding primary aby on all wells, 30min. Dilution buffer & secondary aby dilution buffer is the same (1xpbs/0.5%bsa/0.005%tween). Primary inc. is 2hours, secondary aby inc. is 1 hour. I am using hrp/tmb for 30 min substrate-is that too long? could that be causing oversaturation?
When ODs fall slower than the expected 2-fold dilution made, what is going wrong and at which step?
thank you.
In my experience 30 minutes is too long for an hrp/tmb incubation. Your strong signals will likely start to precipitate by that time and will give you variability. Also, if the incubation isn't happening in the dark, you'll start to get some non-specific background.
As an aside, the standard curve really isn't valid if you change the capture protein between your sample and the std(especially where the standard curve is relying on the affinity of the capture antibody for antibody in the serum and your sample is relying on the affinity of the antibody in the sample for the specific antigen on the plate) . It may be able to tell you there is little antibody versus a lot of antibody but no form of quantitation will be accurate - not even semi-quantitative. It isn't comparing apples to apples - more like apples to carrots 
Good luck!
Sorry, I didn't answer the slow fall in OD part. I don't think it has to do with the length of incubations but rather the amount of antigen on the plate or the starting concentration of antibody in your sample. The slow fall in OD is similar to the plateau effect where there is an excess of antibody. Either your specific protein isn't coating well on the plate (I don't know if you have a control for that or how much you're coating) or there is too much specific antibody in your sample. If you're confident that your specific protein is binding to the plate well then I would try starting the antibody sample at a higher dilution and see if you can get a better signal drop.
As both Missile and I are suggesting, you are running two assays on one plate...it makes little sense to compare the results of your assay with the specific protein coat to a curve with a different coat for the reasons we have both indicated. If you require only relative results...create a pool of serum containing your antibodies for your specific protein, use this to prepare your curve with arbitrary units, and use this as a reference for moving forward.
When ODs fall, what is 'slower than expected'? Each assay format will behave differently depending on the binding affinities and avidities of the antibody/antigen interaction and the curve will fall slower with one format than another. When you say high CVs, do you mean high CVs between replicates of the same sample, or CVs generated from back calculated results at different dilutions of the same sample? If your replicate wells give the same ODs when loaded with the same sample at the same dilution, there is hope for the assay, just stick with the specific protein coat and find a way to standardize the curve so you can compare like with like.
For TMB, as Missile suggests, 30 minutes before stopping TMB is a little on the long side from my experience too.
Good luck
In addition to what Ben Lomond and Missle said (which I totally agree), I have a question. What is your "IgG STD"? I mean, if you are using this as some form of positive standard for the amount of specific IgG in your samples, then (as said before) you should coat the whole plate with your protein, and use that serum (in a dilution series) to make a standard curve. If, on the contrary, the "
Found this in a very quick google search: http://www.elisa-antibody.com/ELISA-applications I'm sure there are plenty more references.
Hope this helps, if this is not what you are trying to do I've completely misread your posts.
Boy is this topic confusing!
Follow the suggestions above
Method/Test1...capture antibody and optimize that test with standards/controls/calibrators and conjugate. Once you get a curve then run your unknows.
method/Test2....Separately, use your binding protein as capture along with standards/controls/calibrators etc etc and do the same thing.
Once you have these 2 tests optimized then results can be compared.
What if you don't have any standards/cntls/calibrators with a known amount of specific aby to the binding protein? How do you establish a STD when you are starting from scratch & the only thing you have to compare to is a commercial STD with a known amount of IgG (not specific). I'm not sure why the protocol was set up the way is was, never made much sense to me.
We have this: 1) a specific protein (I'll call it "X")
2) serum samples for which we are trying to find an amount of IgG specific for our protein
3) a commercial quantitation kit that has a Reference serum with a known amount of IgG (not specific for "X") , an IgG coating antibody, & a secondary IgG hrp-aby
What we DON'T have is this:
a sample to use as a STD/REF curve that has a known amount of aby specific for "X"
I get what Vetetan is saying for Method/Test1...if I coat with our commercial kit of capture IgG & run the Reference serum that comes with it as the STD curve with an assigned conc., that will give me the general IgG conc. for serums that we run on that plate & from that I can assign one of them as a new STD or control with an assigned conc. if I wish
When I get to step 2 & coat the plate with "X", what am I using as the STD for the curve and what conc. do I assign it for the curve? Am I still using the Reference serum from the kit with it's assigned IgG conc.?
sorry this is so confusing, I'm used to establishing assays with some sort of known STD to work with first.