Chromatography Forum

Hi everyone. So I've been trying method development to analyze a mixture of several fatty carboxylic acid derivatives. I am using a Waters Symmetry C18 column, 3x100mm, 3.5um, 100A at a flow rate of 0.5 ml/min, column temp 40C. My HPLC system is a Varian Prostar 335 DAD with 2 210 solvent pumps and a 410 autosampler with 100uL sample loop.

Initially, I tried using plain unbuffered MeOH:H2O. From 70:30 MeOH:H20 to 95:5 MeOH:H2O over 25min. The peak shape of the main component looked nice, however there was an uneven baseline due to gradient drift and several other components did not resolve well. Here is how it looked...



After reading up on some literature, I changed the solvent system to 35mM ammonium formate at pH 5.5. This time the baseline was beautiful, and most of the peaks gave sharp and great definition (some tailing). However the main component now shoulders...



Conclusions I've drawn...
1. Since it is the only peak that shoulders, it is probably not a frit or column guard problem.
2. I adjusted injection volume and concentration assuming overload, but there was not change in peak shape.
3. The injection solvent the sample is dissolved in matches the mobile phase concentration (70:30 MeOH:H2O - but it is not buffered, should it be?).
4. It most likely isn't a band broadening issue, as other peaks would be affected, and the problem occurred after a change in mobile phase.
5. The most obvious reason would be a co-eluted peak, however the purity factor on the DAD revealed the peak was pure, and scanning the UV of the peak across its whole width showed a similar absorbance...



Could I assume the change in peak shape is likely a pH issue? What would you recommend I do next, change buffer concentration and pH (should I go lower for a carboxylic acid?) Try a different column? Any advice would be helpful.

Edit: The buffer solution I made was 3 days old before I had a chance to use it, could this have also been an issue?

Hi

thanks for complete information at first post. not always seen here...

I think your thoughts/conclusions are fine.

By buffering you've got a huge shift in retention, so a pH issue is pretty likely.
Do you know something about your molecule's pKa? Try a buffer pH that is +/-2 units away its pKa, for acids stay lower.

Formate at pH 5.5 isn't the best choice as the pKa of formic acid is 3.7. For 5.5, acetate buffer would fit better (pKa 4.7).
Try to use the buffer within +/- 1 units around its pKa value.

The easiest thing to test would be to try 1 or 2 different buffers, e.g. NH4-formate at pH 4 and 3, or just try plain formic acid 0.1%.

If the retention time is stable even at different pH but still shouldering, try to increase buffer concentration (and/or decrease sample load). How much is the effective injection volume?

And lowering/adjusting the pH of your samples shouldn't hurt as well (for a qualitative test, fill two vials with your sample, to one add some formic acid (ca 10 µL/ml) and compare with the untreated one)

Hello, and thanks for your reply and suggestions. I will try lowering the pH. How high should you go on a buffer concentration before reaching the point of diminishing returns, 50mM?

Yes, I did get a huge shift in retention by buffering. I was under the impression that by lowering the mobile phase pH, acids would show increased retention because ionization would be suppressed? That is why you suggest a pH +/- 2 units from the compounds pka, right? But my compound showed much less retention.

At pH 7, carboxylic acids can undergo hydrogen bonding with active silanol sites leading to tailing. Is there a specific symptom that leads to shouldering?

In addition, the buffered method gave a greater resolution on trace components and overall a better separation. The baseline drift due to the gradient in the first method was also completely resolved by using a buffered aqueous phase.

BTW. Injection volume was between 15-30ul

If your compounds contain anino- groups, they’ll probably be positively charged at low pH and the retention time will decrease dramatically.

Best Regards

Could it be that another compound is eluting very near to your target compound?

I concur with Gergard that it looks an awful lot like a second peak. Peak purity doesn't really tell you much if two compounds have very similar spectra.

Is there any chsnce that your compound may be flipping between two conformers?

Thanks for all the input. I really don't believe it is another compound, there shouldn't be anything else in there that absorbs so strongly. The possibility is always there however. My list of things to try today...

1. Remove guard column and try again.
2. Lower pH of aqueous to 4 then to 3 if needed and try again.
3. Swap out column. I have a new 3x150mm 3.5um C18 waters x-bridge column. It's a bit longer, but a suitable replacement.

Also, I ordered a standard for the compound in question. If it resolves the same way, then I'll know for sure it's not another molecule. Won't arrive until late next week however.

Any other suggestions to try?

Btw, the compound does not have a ring to form a diastereometric pair. Also no amino groups either.

First of all, I'd try an injection of your compound dissolved in the exact starting mobile phase- including the buffer of course, in order to rule out any solvent/mobile phase mismatch issues.
If this doesn't help, try lowering the injection volume drastically, let's say to one tenth, to check for any overloading issues.
BTW if your compound shows less retention with lower pH it must be basic in nature or something's terribly wrong.
What Tom was aiming at is if there was any chance for slowly equillibrating conformational isomers - no need for a ring to have these.

The other possible cause is that the compound was injected as a salt. I have had many similar chromatograms from post purification samples where the answer was to process the sample through an ion exchange column and the analytical problem promptly vanished.

The other possible cause is that the compound was injected as a salt. I have had many similar chromatograms from post purification samples where the answer was to process the sample through an ion exchange column and the analytical problem promptly vanished.

Great, and very insightful suggestion, thank you. Might be a good idea to add the buffered aqueous phase to my sample for injection.

So I've been doing some tinkering. Changed the pH of the aq. buffer to 5.1 and now look at the difference...



The shoulder is definitely gone. Peak still a little mis-shapen, but I'm going to adjust the pH to 4.5-5.0 range and see if that resolves this last little issue. Must be due to a column interaction, probably silanol hydrogen bonding. The 0.4 decrease in pH must have been enough to protonate everything. It always amazes me to see such a small change in pH can have such a dramatic effect.

Thanks for all your input everyone, these boards can be real helpful for chromatographers! Not just for support, but also to validate things you know you should try to do.

It indicates a possible amino- group.
If I were you I'd try even higher pH - like 6.5 phosphate.
The peak is not completely symmetric so I think you maybe too close to a critical pH value.

Best regrads

Update:

Well, finally figured out what was the matter. First off, I replaced my dirty guard column cartridge which decreased the system pressure by almost 1000psi - ! Next, I did a long flush of my column with 95:5 MeOH for about an hour with the new guard... a scattering of junk peaks came out. Finally, I adjusted the pH of my aqueous buffer to 4.50 with more formic acid. Now look at what we have...



Definitely a pH/dirty column/dirty guard issue. Lost a little resolution with the last peak in the above run because the peak eluted right in the middle of a gradient rise.

Danko - once again, no amino groups present in my sample