Chromatography Forum

I think this topic may have been touched upon (as parts of other threads) but I don't recall it being "hit head on".

Let me start by asking in a general sense, what people tend to use for buffers/additives in HILIC methods. Of course, it varies depending on the analyte. But please answer - to the extent possible - in a general sense.

What triggers my question is that we recently tried using triethyl ammonium acetate for a HILIC method (the analyte was EDTA). Not only did it not result in a good peak shape for EDTA, but it caused a pretty bad baseline. It's not really clear to me why this would be the case. And, by the way, we were at 220 nm, so I don't think it should be a UV cuttoff issue causing the unruly baseline.

Thanks for any thoughts.

Once we start considering HILIC as a multi-mode chromatography (where various retention mechanisms are working) it becomes easier to solve problems. As the number of interactions increase for an analyte with the stationary, peak shapes become worse and worse. A decent example is that of chiral compounds where at least three different types of interaction must occur- the result is the chiral separations often have lower efficiency and possibly bad peak shapes. You didn't mention the stationary phase but your analyte is a hexadentate ligand with 4 carboxyl and two amino groups. No wonder the peak shape is bad on a charged stationary phase (usually employed in HILIC). If there are traces of metallic ions from the frits, one can expect worse.

If you read HILIC papers, Alpert (the person who coined this term) recommends a magic 20 mM concentration of buffer concentration to eliminate such electrostatic effects mentioned above. One would normally choose the simplest possible buffer. I can recall using phosphate, methyl phosphonate, ammonium acetate among others. Higher concentration of buffers at neutral pH can very easily dissolve your silica column. Secondly triethylammonium acetate might be changing the surface properties by adsorbing on the surface.

Maybe a bit OT, but here two quick separations of EDTA in HILIC mode.
Gradient from 0 to 100% B in 15 minutes.
A: ACN/Water 8:2, salt 10 mM
B: ACN/Water 1:1, salt 10 mM
flow: 0.2 ml/min
detection: ESI- TOF
Top: Xbridge Amide 150 x 2.1 mm x 2.5 um, salt ammonium bicarbonate not titrated
Bottom: Luna HILIC Diol 150 x 2.1 mm x 3 um, salt ammonium formate not titrated

First peak: Iron EDTA
second peak: free EDTA

Thanks for the publicity regarding the 20 mM salt figure, Farooq. However, that pertains to the easy cases where you wish to disrupt electrostatic effects between the stationary phase and an analyte with some charge but not a lot of it. EDTA is a special case. Whenever you have an analyte with numerous ionizable groups and a high charge-to-mass ratio like this, then you need even more salt to insure that all of the ionizable functional groups share the same counterion. Otherwise, different charged group-counterion pairs will differ in polarity and will migrate at different rates in HILIC. Carlo shows us a nice example of this with EDTA with and without chelated iron. I examined this issue in some detail in my 2008 paper that introduced the ERLIC mode, cf. the following link: http://pubs.acs.org/doi/pdf/10.1021/ac070997p
Specifically, look at Fig. 14, which concerns the migration of arginine. If the counterion in the sample solvent doesn't match the counterion in the mobile phase, then arginine elutes in two well-separated peaks with a continuum between them. One peak corresponds to arginine with the original counterion, the other peak corresponds to arginine molecules that exchanged their counterion for the one in the mobile phase at the beginning of the run, and the continuum corresponds to arginine molecules that exchanged their counterions sometime during the run. A less extreme case would result in a badly skewed peak. The take-home lesson is that for analytes like EDTA with an extreme charge-to-mass ratio, a salt concentration of at least 40 mM is required in the mobile phase and in the sample solvent. In even more extreme cases, such as aminoglycoside antibiotics (kanamycin, etc.), you need at least 100 mM salt to get symmetrical peaks in the HILIC mode.

Interesting feedback. Thanks all.

I would like to ask a related question, while we're talking about this. When developing a HILIC method, do you generally endeavor to get the analyte in the ionized state (and I ask this not for EDTA, but in general). I have heard people say that it is desirable, when doing HILIC, to get the analyte in the ionized state: the reason being simply that in HILIC this gives us more retention (hence, hopefully, more resolution) which is quite logical.

But recently I've been doing a fair amount of reading about acid/base effects in non-aqueous systems, and I am starting to doubt that the above strategy is realistic. Consider a hypothetical basic molecule with a pKa of 5. So, in an aqueous environment (or at least mostly aqeuous) one would set the pH to maybe 3.5 in order to ionize the molecule. But if we now think about a typical HILIC starting mobile phase of 95% ACN/5% Water, everything changes -- and pretty dramatically. First, we need more acid to reach this same pH, but even if we added sufficient acid, it still wouldn't do the job because the pka of the analyte would also have shifted down several pH units. Bottom line: it seems to me that, in most cases, we would need an awful lot of acid to get basic analytes protonated in a 95% ACN mobile phase (and likewise, an awful lot of base for acidic molecules). And on top of all this, as Andy points out above, even if we could accomplish this the analyte would most likely exist an ion pair, so it's still not really charged. So the notion that we can get the molecule to be highly retained by virtue of ionizing it just seems unrealistic to me.

Any thoughts. Please share if you think there's something I am missing.

I would like to ask a related question, while we're talking about this. When developing a HILIC method, do you generally endeavor to get the analyte in the ionized state (and I ask this not for EDTA, but in general). I have heard people say that it is desirable, when doing HILIC, to get the analyte in the ionized state: the reason being simply that in HILIC this gives us more retention (hence, hopefully, more resolution) which is quite logical.

That is a very interesting question but perhaps very difficult to answer. However, the key point lies in my first reply: HILIC is a multi-mode chromatography (forget about the simplified picture of partitioning of the analyte in stagnant water layer for the time being). Ion-exchange is of the retention mechanisms in HILIC. If someone wants to employ this mode, you need ionized analytes.

You need to know what is on the stationary phase to make sensible choice of the pH. BTW, Dr. Alpert recommended 100 mM buffers for complicated cases. Just keep in mind that polymer coated silicas (such as those made by Dr. Alpert) may handle such buffers but hydrophilic bonded phases will simply say good-bye to the silica surface within a day or so. Many bare silicas will also dissolve with such buffers if the organic content is lower than 80%.

Coming back to the question of shifts in pKa of the analyte. Many chromatographers are unaware of the concept of the pH of the stationary phase (again you need to know what is the surface chemistry of your HILIC phase). This point was highlighted by a highly experienced gentleman. I leave the following as a gedanken experiment:

Imagine you have 1 mM NaOH as a mobile phase being used on a anion-exchanger. The mobile phase pH is 11. However on the ion-exchanger, all sites are saturated with OH- as the counter-ion. What is the stationary phase pH? If the analyte comes in contact with such a stationary phase, what would be the state of the ionization of the analyte?

Regards,

Farooq

I have a question on this topic.

I have recently heard that equilibration in HILIC can be very slow when we have a mobile phase of 95% ACN and 5% water. And that it is much quicker at 10% water or especially at 15% water.

This surprised me, because it seems like a lot of HILIC work is done at 5% water - or at least the gradient starts there.

So I want to ask, what is the reason that equilibration is slow when there's only 5% water in the mobile phase?

Thank You
Adam

I have recently heard that equilibration in HILIC can be very slow when we have a mobile phase of 95% ACN and 5% water. And that it is much quicker at 10% water or especially at 15% water.
This surprised me, because it seems like a lot of HILIC work is done at 5% water - or at least the gradient starts there. So I want to ask, what is the reason that equilibration is slow when there's only 5% water in the mobile phase?

I am not sure if 5% is a cut-off or specific number but the general trend is true. HILIC is like normal phase, with the difference that the solvent is miscible with water in all proportions. For instance in the normal phase, water is present in a very small quantity in solvents like hexane and in ethanol. It often takes a long time to establish an equilibrium to rehydrate the silica surface with these solvents. Similarly, in the HILIC mode when the concentration of water is low in the mobile phase, it may take a while to establish a water layer in equilibrium in the polar stationary phase.

Many HILIC phases have polymeric coatings. With the changes in mobile phase polarity there might be reorientation effects of the polar groups and hydrocarbon chains. The kinetics of this process might be slow as compared to gradient speed.

That's interesting. I know that if the water percentage is too high (like maybe over 70%) then you don't form the sorbed water layer at all, basically because the water molecules are "happier" in the mobile phase as opposed to sorbed to the surface. So it's a thermodynamic issue. But what you're saying is that if the water content is on the low side (say around 5%) then, although the process is quite favorable thermodynamically, there is a kinetic issue. So it takes a while.

I guess the idea is somewhere in between these kinetic and thermodynamic extremes.

I wonder if it might be a recommended practice, in a case where we are running a gradient from 5% water to 50% water, to re-equilibrate the column at 15% water for 5 minutes (because of the much faster kinetics), and then drop to 5% for another 5 minutes (or something along those lines), so that the equilibration can be done more quickly, rather than requiring a 30 or 40 minutes equilibration after each run. Anyone ever try this.

I haven't tried that, but when I want to speed up the admittedly lengthy approach to equilibrium with low percentages of water, I double the flow rate. This cuts the equilibration time in half. Backpressures are generally quite low with such high percentages of ACN, so you can get away with this.

That's interesting. I know that if the water percentage is too high (like maybe over 70%) then you don't form the sorbed water layer at all, basically because the water molecules are "happier" in the mobile phase as opposed to sorbed to the surface. So it's a thermodynamic issue. But what you're saying is that if the water content is on the low side (say around 5%) then, although the process is quite favorable thermodynamically, there is a kinetic issue. So it takes a while.

Dear Adam, Although it may sound counter-intuitive, but the more water you have in the mobile phase, the more gets adsorbed on the stationary phase on any polar surface. So the hypothesis of assuming that a water layer is not formed in high water content mobile phases is not correct. The reason for early elution with high water content is that the mobile phase is very strong. It is just like an adsorption isotherm experiment (the adsorbate is water in this case): the more amount of "x" you add, the more "x" gets adsorbed on an adsorbing surface, till it reaches a saturation value like a Langmuirian curve. HILIC mode is no exception. When water concentration is low, it takes a while to reach the equilibrium value in a given mobile phase.

Isn't the thread going off-topic? I must admit it's partly my fault. The initial question was about use of buffers/additives. So I will just put forth my simple answer: I think TEA/acetate buffer does not work fine for HILIC because TEA is an ion pair reagent. As an example, on a Waters XBridge amide If I use ammonium bicarbonate pH 9.5 I can see at the same time sugars and organic acids; while if I use TEA i will only see the sugars (all the acids elute very early and with no separation)