Can you report analyte content without running standards eve

Basic questions from students; resources for projects and reports.

6 posts Page 1 of 1
Hi, I'm wondering how you could or if you could report the quantity of an analyte if you do not run standards with a known concentration.

Right now I'm working on a project to examine the various terpenes found in hops. I have about 15 terpenes of interest. I have standards that I have run to verify retention times, but each standard is expensive, and it just wouldn't be possible with the budget the school allots to run the standards every day that I test new samples. To buy the individual 1mg/ml standards It cost me about $1000. I made a working stock, but if I run a calibration curve everyday I run samples than my working stock only lasts three days.So if I were to have to keep buying standards and even after diluting them for a working stock, Id essentially to spending around $1000 every three days by the end of my project I would have to spend close to $40,000 in standards alone.

Is there anyway to calculate the content from the area of the peak, or would I just report the relative percentages of the terpenes found in the hops being tested? I'd love to get some insight in what to do in situations like this.
In principle, "no" (chromatography is always a *relative* technique for quantitation). In practice, "maybe".

The answer hinges on the purpose and behavior of the analysis.

If this is a regulatory assay (labelling for human consumption), then the answer would be a definite "no".

Otherwise, you have to look at how consistent the response factors (RFs; slopes of the calibration plots) are over time. Go back and look at your history and see how much variability there is on a day-to-day basis (also check to see if there is any trend). If that level of variability is acceptable for your purpose, then calibrate less frequently.

If the variability is more than you can tolerate, then look at the relative response factors (RRFs). If those are consistent, then you should be able to use an inexpensive internal standard to do your daily calibration.

If there is too much variability in the RRFs, then you're back to "no".
-- Tom Jupille
LC Resources / Separation Science Associates
tjupille@lcresources.com
+ 1 (925) 297-5374
Much of the cost of the standards you are using goes into the time needed to do the dilution (and certify it if you are working at that level). Costs might be substantially reduced if you by the pure substances and dilute them yourself.

If you have a method that works well, with little discrimination between analytes (which is entirely possible with terpenes) then you might do better with internal standards rather than running a full external calibration with every batch.

Peter
Peter Apps
If this is a research project with no regulatory implications, then there are all sorts of lesser options you can consider:

Of course Peter is right, bulk compounds are cheaper than deliberately prepared standards, but for many compounds you'll struggle to get them cheaply in any form. If you can find a closely related compound that is available more cheaply, you can use this as a standard, and express your results on that basis; it will, at least, give some level of standardisation from experiment to experiment, and reduce variation caused by instrument efficiency. In some fields there is no choice but to use a related, non-identical standard: for example, the plant kingdom contains well over 100 different glucosinolates, but so far as I know only one is available as a commercial compound, so there are an awful lot of publications that estimate all their glucosinolates based on sinigrin. This isn't evil, provided you make it clear what you are doing. It's slightly annoying having to do it, because most chromatography software will, with justification, dislike cooperating! You would need to run your standard curve using cheap-X, then apply it to a set of peak areas at different retention times, which quite probably means doing the area-to-amount calibration curve bit outside the chromatography software - always an unhappy thing to do. You can also consider measuring the relative response factor of the cheap-X standard compared to the expensive standards of compounds A, B, C... and doing a conversion by using this "fudge factor" each time. Again, this isn't evil, provided you state what you're doing, and consider all possible ways in which it could go wrong.

Big thing: if your calibration curves are always linear, then you don't need to prepare a whole dilution series each time. Consider one-point calibration, and don't replicate your curve more than you can afford.

Consider for each experiment whether you are interested in absolute amounts, or looking for changes. If you're looking to see if a variety of hops is different, or if you're screening loads of lines of hops hoping to find one that's different, you don't need to estimate the absolute amount, and raw peak areas are quite adequate. Only perform the calibration when there is a real need for calibrated, absolute results.
lmh wrote:
If this is a research project with no regulatory implications, then there are all sorts of lesser options you can consider:

Of course Peter is right, bulk compounds are cheaper than deliberately prepared standards, but for many compounds you'll struggle to get them cheaply in any form. If you can find a closely related compound that is available more cheaply, you can use this as a standard, and express your results on that basis; it will, at least, give some level of standardisation from experiment to experiment, and reduce variation caused by instrument efficiency. In some fields there is no choice but to use a related, non-identical standard: for example, the plant kingdom contains well over 100 different glucosinolates, but so far as I know only one is available as a commercial compound, so there are an awful lot of publications that estimate all their glucosinolates based on sinigrin. This isn't evil, provided you make it clear what you are doing. It's slightly annoying having to do it, because most chromatography software will, with justification, dislike cooperating! You would need to run your standard curve using cheap-X, then apply it to a set of peak areas at different retention times, which quite probably means doing the area-to-amount calibration curve bit outside the chromatography software - always an unhappy thing to do. You can also consider measuring the relative response factor of the cheap-X standard compared to the expensive standards of compounds A, B, C... and doing a conversion by using this "fudge factor" each time. Again, this isn't evil, provided you state what you're doing, and consider all possible ways in which it could go wrong.

Big thing: if your calibration curves are always linear, then you don't need to prepare a whole dilution series each time. Consider one-point calibration, and don't replicate your curve more than you can afford.

Consider for each experiment whether you are interested in absolute amounts, or looking for changes. If you're looking to see if a variety of hops is different, or if you're screening loads of lines of hops hoping to find one that's different, you don't need to estimate the absolute amount, and raw peak areas are quite adequate. Only perform the calibration when there is a real need for calibrated, absolute results.


I work with glucosinolate analysis and I can confirm they are expensive for standards, but there are several that are available just be prepared to spend something like $500 for 10mg or more of each one. We look for Glucoraphanin on a regular basis and if you want accurate numbers you do have to use it to calibrate because Sinigrin is not 1 to 1 on response versus concentration.

For the terpenes analysis, you could try what we do in the environmental lab, where you calibrate once, then check the calibration each day with one level of standard, if it passes with say +/-10% or 15% then call your calibration good and use it that day for analysis.

How much working stock volume are you making and why does it only last three days? I would think it would last longer that that without going bad. If you recap the vials and store in a cooler it should last quite a while unless they break down really fast.
The past is there to guide us into the future, not to dwell in.
ntward,

So my other questions is..... On your other post you say you are calibrated at ng/mL and you are buying mg/mL so you should have a lot of room here to make dilutions, right? If so, then a daily standard of all components should be available from the dilution. Sorry to ask but I have to, is your ng/mL (ppb?) actually what your standards are at?

Best regards,

AICMM
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