In the previous instalment of HPLC Solutions (HPLC Solutions #134) we took a brief look at why we might decide to use an internal standard and how a calibration plot is constructed and used in HPLC analysis. This time, we’ll consider some properties that are necessary when choosing a compound to use as an internal standard. These are listed in the figure.
In the next few instalments of HPLC Solutions , we’ll look at several aspects of the use of internal standards (IS) in HPLC and LC-MS applications. The topic of this discussion is how an internal standard is used in quantitative analysis.
This is the fourth instalment of HPLC Solutions related to resources you should have at your fingertips when you are working in an HPLC laboratory (see also #130, 131,132). In the last discussion (#132) we looked at online resources from the International Conference on Harmonization (ICH) and the United States Food and Drug Administration (FDA) regarding validation of HPLC methods. This time I’d like to mention a few books that I find indispensable in my day-to-day work as a chromatographer. The first three form my trifecta of HPLC books that I refer to most often.
This is the third instalment of HPLC Solutions related to resources you should have at your fingertips when you are working in an HPLC laboratory (see also #130, 131). If you work in a regulated industry, such as pharmaceuticals, your HPLC methods likely will need to be validated. If you are using a compendial method from the United States Pharmacopoeia (USP) or European Pharmacopoeia (EP) or other pharmacopoeial method, you may not have to validate the method, but simply demonstrate that it passes system suitability. If the method is modified sufficiently in transfer, it will need some sort of validation.
In this instalment of HPLC solutions , I’d like to share a few of the places that I look when confronted with a new separation problem.
I’m sure you’ve all heard the proverb that says if you give a person a fish, you’ll feed them for a day, but if you teach a person to fish, you’ll enable them to feed themselves. There are lots of on-line options for “fishing lessons” through LC Resources and Separation Science, many with free access. In the next few HPLC Solutions instalments, we’ll look at other resources focused on specific tasks you need to undertake.
In the first two parts of this series (HPLC Solutions #127 and #128), we looked at how HPLC data systems integrate chromatograms and some of the adjustments that can be made if you don’t like the way the default settings perform. Even with proper adjustment of the settings, you may still observe occasional or regular problems with integration. Here we’ll look at several common problems and how to correct them.
In HPLC Solutions #127 we took a brief look at how HPLC integrators detect and measure a peak’s height or area. We’ll look here at some of the options available to adjust the integration process.
With today’s data systems for HPLC, we tend to take peak measurement for granted. Much of the time we’re pretty safe if we use the default integration conditions, but this is not always the case. In this HPLC Solutions series on integration, we’ll look at how peaks are integrated and what can go wrong. This instalment centres on the integration process itself.
HPLC Solutions #126: Chromatographic Measurements, Part 5: Determining LOD and LOQ Based on the Calibration Curve
In this issue of HPLC Solutions we look at determining LOD and LOQ based on the calibration curve.
HPLC Solutions #125: Chromatographic Measurements, Part 4: Determining LOD and LOQ Visually or by S/N
In this issue of HPLC Solutions we’ll look at the first two methods, and then consider the third in #126. For additional references, you might want to look at #122 for the determination of noise and #124 for calculation of the signal-to-noise ratio (S/N).
In HPLC Solutions #122, we looked at how to measure baseline noise from a liquid chromatogram. One practical application of such measurements is to determine the signal-to-noise ratio (S/N), which we’ll consider here.
In HPLC Solutions #122, we looked at how to measure baseline noise from a liquid chromatogram. One practical application of such measurements is to determine the noise and drift of the detector, which we’ll consider now.
This installment of HPLC Solutions begins a series on different measurements that we make on chromatograms. Measuring the baseline noise generated by an HPLC system may seem like a singularly boring subject. But if you think about it, noise can be a critical factor in the results of our analytical work. It is directly related to the limit of detection and limit of quantification, it can be used to determine the health of the detector, noise is a great diagnostic tool for problems, and can be a limiting factor in the precision of a method.
In the last instalment of HPLC Solutions (#120) we looked at one popular autosampler design, the push-to-fill autosampler. Now we’ll turn our attention to the other most popular design, which I call needle-in-loop.
As a follow-up on the last two HPLC Solutions discussions (#118, #119) regarding sample injection, I’d like to take the opportunity to describe the two most popular types of autosamplers in use today. These are what I call the “push-to-fill” design that is discussed here, and next time we’ll look at the “needle-in-loop” design.
This discussion follows on the last HPLC Solutions instalment (#118), where we discussed the relative merits of manual injectors and autosamplers. A little better understanding of the loop filling characteristics will help us realize why injection volumes aren’t always accurate and why autosamplers tend to be more precise than manual injection.
Q: I’m wondering why most of scientists prefer to use a manual injector?! I know that autosamplers are used more to improve the productivity and it is standard with many HPLC systems. I was working with both and the autosampler produced better precision. Please can you tell me in which situations one is better than the other?
Most of us take advantage of the autozero function on our HPLC data systems to make the chromatogram look more pleasing to the eye. The autozero function is quite simple – it just takes whatever signal the data system is receiving and sets it at the zero point on the y-axis of the output.
Q: I have just developed a gradient HPLC method and need to set the acceptance criteria for system suitability. The column efficiency is one of the measurements we usually include in system suitability, but I don’t know which peak I should use to measure the efficiency. Can you help me?
A question that commonly arises in my Master Classes is the proper technique to integrate chromatograms where the peaks are not fully baseline resolved. For this, I most commonly draw on what I call the 10% Rule. This applies when a minor peak is partially merged with a major one. If the height of the minor peak is <10% of the height of the major peak, a skim is the appropriate integration technique. If the minor peak is >10% of the major peak, a perpendicular drop is the best technique.
For the last few instalments of HPLC Solutions, we’ve been looking at ways to reduce baseline noise. We’ve been particularly focused on noise from electrical sources which tend to reflect the frequency of the power delivered to the laboratory.
In the last instalment of HPLC Solutions (#112), we looked at one of the two most common ways to filter the detector signal to reduce baseline noise – the detector time constant. The second technique is to adjust the data system data rate.
In the last HPLC Solutions (#111) we looked at the influence of signal averaging on baseline noise for a simulated chromatogram. Let’s look now at how the detector time constant can be used to reduce baseline noise.
In HPLC Solutions #110 we looked at a problem of baseline noise created by a pump malfunction. In this instalment, we’ll look at what can be done about electronic noise in the chromatogram.
In HPLC Solutions #109 we discussed a technique to measure noise and drift for an HPLC detector. There are several possible sources of noise in the detector, and the approach to reducing or eliminating such noise depends on the noise source. For the next few HPLC Solutions installments, we’ll look at some different sources of noise.
One of the topics in my HPLC Troubleshooting: A Performance Qualification Approach Master Class is the importance of knowing how your HPLC system works when it is working correctly. Knowing the normal detector noise and drift can help you to realize when there is a problem, by giving you a reference value for your detector in addition to the detector specifications provided by the manufacturer.
A reader observed baseline drift in his gradient method and wondered where it came from and what could be done to eliminate it. There are two main sources of baseline drift – temperature and mobile phase. If the column is operated in a column oven, the temperature should be constant, so temperature-related drift usually is not a problem, since most HPLC systems are configured with ovens.
A reader recently asked how to determine the column volume of a column packed with superficially porous particles. You’ll recall that these are a newer type of HPLC packing material, variously referred to as superficially porous, core-shell, fused-core, shell-type, pellicular, or some other name to describe the fact that the particle comprises a solid center (core) covered by a porous layer of silica, where the chromatography takes place.
A reader asks, “I am using a shared equipment where somebody used tetrabutyl ammonium (TBA) as an ion pair in their mobile phase for chromatography. It looks to me like the ion pairing reagent was not washed completely from the column, even though I flushed repeatedly with methanol (MeOH). Is there a way I can get rid of it?
A reader asks, I would like to split HPLC column effluent between an evaporative light scattering detector (ELSD) and a fraction collector. What is the best way of doing this, particularly as we may want a biased split, i.e., 10% to detector and 90% to fraction collector?”
A reader recently submitted this question: “Most HPLC methods call for preparing the sample in mobile phase as the solvent. I like to avoid using some of the organic solvents for samplepreparation because of toxicity issues, so most of the time I will prepare the solutions using only the aqueous component of the mobile phase as the sample solvent if the analyte is sufficiently soluble in water. For example if the mobile is 80/20 acetonitrile/buffer, I will use the buffer as the sample solvent. Am I creating a problem when I do this?”
A reader recently asked, “The flow cell on my UV detector is completely blocked with some kind of white substance. I tried using a syringe to flush with water and with isopropanol, but neither was successful (and the process was very slow). The manufacturer of the detector says that I can get a cell service kit, but I’m a graduate student and can’t afford one. Do you have any ideas what is going on and how to fix the problem?”
John Dolan relates the tale of how one of his Master Class students left instructions with her lab mate to shut down her HPLC system whilst she went away on vacation for two weeks only to return to find that her friend had only turned the power off.
A question that comes up now and then in our Master Classes is how to figure out the volume of an HPLC column. I’d like to share a couple of rules of thumb that I find very useful for estimating column volume, VM
How do you go about recovering a column that you suspect has some problems?
What do you do if you receive the gift of an HPLC system of unknown breeding?
A reader is trying to figure out the fate of a pesticide in water.
A reader asks, “I have a nice isocratic method for compound A, which elutes at about 8 min, so I can inject a new sample every 10 min. Occasionally, however, my samples contain another compound B that comes out at about 40 min. I have no way of knowing in advance if B will be present or not"
A reader asked me when it is appropriate or necessary to premix mobile phases when on-line mixing is available.
I recently had an “Ask the Doctor” question that went something like this: “I need to report impurities for my product at levels of 0.1-0.5% of the main ingredient. Occasionally when I show my folks my chromatograms, I’m accused of making the sample too concentrated, because the product peak is >2000 mAU. They tell me that the detector is saturated, so the area-% for the impurities will not be accurate. Can you comment on this?”
In a previous article (#93) we looked at a problem gradient method where peaks in the second and subsequent chromatograms were eluted too early. Our conclusion was that the most likely cause was insufficient equilibration time between runs.
A reader emailed in the following question: I am trying to analyze a peptide (2675 Da) on a 150 x 2.1 mm, 3 µm particle column by LC-MS using an ion trap MS in the positive electrospray ionization (ESI) mode. The first injection is OK, with the peptide eluting at ≈ 15 min with good peak shape, but in subsequent runs the retention time is ≈ 5 min. If I try to run the method the next day, the same pattern occurs. The A-solvent is 10% acetonitrile (ACN) in 0.1% trifluoroacetic acid (TFA); B is 100% ACN with 0.1% TFA added. The gradient runs from 0-100% B over 30 min, then steps back to 100% A for 20 min prior to the next injection. Can you spot any obvious problems here?
In a previous article we looked at a problem where we suspected that the data acquisition rate of the data system was too slow to gather a sufficient number of data points across a peak with a 2-µm particle column. This article looks at those calculations in a little more detail, in case you want to perform a similar exercise for your HPLC system.
I recently had a reader send me an email complaining that the peaks in his chromatogram were too narrow. At first, this seems a bit odd – after all, most people complain because peaks are too wide, so narrow peaks usually are preferred over broader ones. Digging into the question a bit more will highlight the real problem.
I recently had an email from a reader who wondered how it was possible to use an evaporative light scattering detector (ELSD) with an HPLC for quantitative analysis when instead of a linear response, the concentration vs response relationship behaves more closely to a power curve.
This is the second article in a series about robustness testing for HPLC analytical methods. Previously, an overview of the steps involved in investigation of robustness was described. The first step is to decide which factors are going to be studied. In this article the selection of appropriate robustness factors and factor levels relating to the mobile phase will be discussed and in particular, the volume fraction of the organic solvent in the mobile phase for reversed phase HPLC.
In an earlier article (#88) we looked at the need to use a matrix-based calibration curve so as to correct for low recovery when extracting an analyte from a sample matrix. To do this, we used a blank matrix and spiked in known concentrations of reference standard to create the calibration samples. But what happens if a blank matrix is not available?
Recently a reader sent me a question regarding a problem he was having with recovery of a vitamin he was measuring in pet foods. He found that the precision, measured as repeatability of peak area for a given sample, was adequate for multiple products, but the assay amount was low by 10–40%, depending on the product. After further probing of the conditions, I found that he was running a calibration curve based on reference standards dissolved in an aqueous solution, instead of spiked into blank sample matrix. I feel that this is the basis of the problem.
In a previous article (Measuring Dwell) we looked at a simple technique to measure the dwell volume of an HPLC system. In a prior discussion (Dwell Differences), it was seen that differences in dwell volume between two HPLC systems could result in offset retention times as well as possible changes in resolution for early-eluted peaks. Another potential problem can be diagnosed using the same dwell-volume test, as is discussed below.
In a previous article (Dwell Differences) we saw an example of how the appearance of a chromatogram could change when a gradient method was run on two HPLC systems of differing dwell volume. In this article we’ll look at a simple way to measure dwell volume.
The dwell volume comprises all the HPLC system volume between the point the solvents are mixed to the head of the column. For high-pressure-mixing systems, this includes the mixer, connecting tubing, and sample loop. For low-pressure-mixing systems, we have the same components plus the volume of the pump heads and check valves. In this article let’s look at some of the problems that can be created when the dwell volume differs between two HPLC system.
A reader recently asked a question about the use of the peak-purity function of his diode-array UV detector (DAD). The question related to whether or not he could detect the presence of enantiomers, stereoisomers, diastereoisomers, or epimers with the peak-purity function.
A reader recently emailed me a question that went something like this: “I’m using LC-MS/MS to analyse a biomolecule that I have isolated from tissue by LC-MS/MS. The lowest point on the calibration curve that I prepared is 0.1 µM. All the samples we’ve tested have concentrations above 0.1 µM and the signal-to-noise (S/N) is 33 at this concentration. I know that S/N = 10 is considered the lower limit of quantification (LLOQ) – does this mean that I have to extend my curve downward, even if I’ll never see concentrations that low?”
A reader reported a problem with blockage of a column packed with one of the newer shell-type packings. The mobile phase A comprises a mixture of 0.1 M KH2PO4, 0.05% triethylamine (TEA), and 150 mg/L EDTA adjusted to pH 7.5 with NaOH. The B-solvent is a mixture of acetonitrile (ACN) and methanol (MeOH). A gradient of 5-60% B is run over one hour, then the column is flushed with 5% B. When he switched from 60% B to 5% B, the column became blocked. Continued attempts at flushing or back-flushing didn’t help. He asked what can be done to rescue the column and to avoid the problem in the future.
After I wrote an article on in-line degassers (In-line Degasser), I immediately received some reader feedback on a specific failure, which was published in 'Failed Degasser'. Since that time I have been collecting occasional reports of degasser failures. This week I’d like to share a few of these.
What happens if your favourite HPLC column accidentally rolls off the bench and lands on the concrete floor of the lab? Is it ruined?
A reader asked me if she was in danger of shortening the lifetime of an HPLC column by removing it after each batch of samples and storing it. This also raised the related question about expectations of shorter column lifetimes if the same instrument was used for multiple methods, each with a different column.
I recently had an inquiry from a reader of 'HPLC Solutions' that comes up with some regularity in the HPLC troubleshooting classes that I teach: “Will I damage the column by getting air in it?” Let’s consider the possibilities for a moment. One possible source of air is injection from an empty vial. Another source is forgetting to put the plugs in the ends of the column when storing it. A third possibility is that you pump the column dry if you run out of mobile phase. What happens in each of these cases?
A reader wrote me recently asking what would happen if he injected his sample, which came dissolved in hexane, into a reversed-phase mobile phase of methanol-buffer. The first answer is that this is not a good habit, but is it possible? We’ll see.
A reader wrote to ask why he was seeing such a difference in the appearance of a gradient chromatogram when all he did was change the flow rate. He’d been making similar changes with isocratic methods for years – an increase in the flow rate shortened the run time, but had little other impact on the separation. Let’s take a look at what is going on here.
In the last instalment of HPLC Solutions (#74), we looked at the gradient proportioning valve (GPV) test as a tool to check for proper operation of the proportioning valves used to blend solvents in low-pressure-mixing HPLC systems. The test comprised steps between water and water-acetone mixtures, testing all possible combinations for the four (A, B, C, and D) solvents. If the steps or “peaks” in the resulting plot are within 5% of each other, the system is operating properly – usually the maximum difference between steps will be 1-2%. Let’s look now at an example where something goes wrong.
If you have a low-pressure-mixing system on your HPLC, the solvents are blended using a proportioning valve. Usually this mixes up to 4 different mobile phase components. The pump delivers at a constant flow rate, for example, 1 mL/min, and the various proportioning valves open and close to allow the right proportion of each solvent to enter the mixer. For example, if you have programmed the HPLC to deliver 75% buffer (the A-solvent) and 25% methanol (MeOH, the B-solvent), the valves would open A-B-A-B-A-B… in a 75:25 ratio of opening times. These pulses of A and B are mixed, drawn into the pump, and delivered to the column. To deliver a gradient, the ratio of the opening times of the various proportioning valves will change during the gradient to change the mobile phase composition.
In the previous issue (HPLC Solutions #72) we looked at how in-line degassers work. This week I’d like to look at an interesting case study related to the degasser that was contributed by a reader (WLT). The story goes like this: I am running a gradient from 0 to 95% B over 10 min on an HPLC system with low-pressure mixing (mixer before the pump). There is a significant amount of retention time drift (0.1-0.2 min, Figure 1) for all peaks. There is no trend of increasing or decreasing retention time from run to run. The service engineer has been in to check for leaks and even changed the multi-channel gradient valve but the problem has not been resolved.
For the first part of my HPLC troubleshooting career, it was very easy to name the number one HPLC problem – degassing. Air bubbles in the pump were a headache for nearly every user. Helium sparging was the gold standard in degassing, but was inconvenient and expensive – and without proper venting, could put unwanted solvent vapors in the laboratory air. Vacuum degassing worked fairly well and when combined with sonication, was effective for most applications. But degassing was always a bother and even with good lab habits, bubble problems were still number one on the problem list. Then came the in-line vacuum degasser. This changed everything and most new HPLC systems come with an in-line degasser. Their effectiveness is reflected in my observation that I hardly ever hear a complaint about air bubbles in the pump anymore. Let’s take a moment and see how these devices work.
One question I get regularly in the classes I teach regards the new porous-layer bead packing materials. Are these for real or are they just hype? Let’s take a look at some of the characteristics of these column packings and see what lies between the frits of the column.
I had a follow-up inquiry about one of the recent HPLC Solutions articles. In Solutions #64 (Why Acid?), I discussed some of the reasons why acids such as formic or trifluoroacetic (TFA) are added to mobile phases for LC-MS work. The reader noted that in Figure 1 of that article, I suggested putting the acid in both the aqueous A-solvent and the organic B-solvent. She said that she only put it in A so that she could have pure ACN in B and then could use it with other mobile-phase combinations as well. Is there anything the matter with this?
Q: I am running an isocratic method with a 250 x 4.6 mm, 10 µm particle column at 1 mL/min with a 20% acetonitrile (ACN)/buffer mobile phase. I dissolve my sample in ACN and inject. Normally my compound of interest comes out at 7.7 min. I switched to a 250 x 4.0 mm column, keeping everything else the same, but now I see a distinct hump on the front of the peak. Is this another compound or am I doing something wrong?
In a recent Ask the Doctor email, a reader enquired about the best way to clean the frits used in the HPLC system. He indicated that he had been sonicating them in nitric acid, but wasn’t sure if that was the best way to approach the problem.
A reader complained about retention times that changed in an overnight HPLC run and wondered how to diagnose the problem source. The question left too many possibilities, and because there was no complaint about changing resolution, I’ll assume that all the peaks moved in the same direction to longer or shorter retention times.
Recently, a reader sent a question to our “Ask the Doctor” e-mail asking how to get started with a method that required separation of enantiomers by reversed-phase HPLC. Unfortunately, the problem is a non-starter in this context. Enantiomers cannot be separated by reversed-phase techniques – they require some chirality in the system.
A reader recently sent in this e-mail: “I am trying to find information that I can put in an SOP regarding how long a mobile phase can be used before it must be discarded. I’ve heard 1-2 weeks from some sources, whereas others say a month, and some people say not to worry about it – just use the mobile phase until the bottle is empty. Can you shed some light on this matter?”
Recently, I received an “Ask the Doctor” email from a reader asking why formic acid was specified as an additive for the mobile phase in an HPLC method he was using. Formic or trifluoroacetic acid at 0.1% concentrations are common, especially for LC-MS work. There are a number of reasons for adding an acid at low concentration to the mobile phase. Let’s look at two of these: the influence on the column and the sample.
PEEK (poly-ether-ether-ketone) tubing and fittings have become a standard part of most HPLC systems today. The flexibility and ease of cutting the tubing, coupled with the convenience of finger-tightened fittings make a combination that is hard to beat for most applications. In at least two of the previous HPLC Solutions discussions (#9 and #55), I’ve discussed various aspects of PEEK products used in the HPLC environment.
In the past instalments of HPLC Solutions (#56-61), we have explored the concept of the adjustment of HPLC methods. In particular, we have tried to determine the difference between method adjustment and method change. The reason for this splitting of hairs is the interpretation of regulatory guidelines that adjustments can be made to meet system suitability requirements, but changes will require some level of revalidation. As a reference point, we used the European Pharmacopoeia (EP ) and United States Pharmacopoeia (USP ) recommendations. The problem with any discussion of method alteration is that there is a lot of interpretation that goes on.
This is the last in a six-part series on allowed adjustments to HPLC methods. In the previous instalments (HPLC Solutions #56-60), we’ve looked at each of the specific variables listed in Table 1.
HPLC Solutions #60: Method Adjustment vs Change Part 5: Volume, Column Temperature and Detector Wavelength
In the previous few articles, HPLC Solutions has focused on the guidelines for method adjustment listed in the European Pharmacopoeia (EP ) and United States Pharmacopoeia (USP ). We looked at pH (#56), mobile-phase composition (#57), column dimensions (#58), and flow rate and particle size (#59). This week we’ll focus on the remaining variables listed in Table 1.
In prior instalments of this series on method adjustment, we’ve looked at the guidelines from the European Pharmacopoeia (EP ) and United States Pharmacopoeia (USP ) for allowable adjustments in pH (HPLC Solutions #56 ), mobile phase (#57), and column dimensions (#58). This week we’ll look at flow-rate and particle-size adjustments.
In this series on method adjustment vs method change, we first took an overview of the concept (HPLC Solutions #56 ) then at specific guidelines for mobile-phase adjustment (#57). In this article, we’ll take a closer look at some of the potential adjustments related to the column.
In the previous instalment (HPLC Solutions #56), we took an overview of the concept of method adjustment vs method change in reversed-phase HPLC, in light of the guidelines of the European Pharmacopoeia (EP ) and United States Pharmacopoeia (USP ). We also looked at an example of the allowed changes in mobile phase pH and buffer strength. In this article, we’ll move down the chart and look at the variables listed in Figure 1.
We all know that no HPLC method is perfect and that it may need to be “tweaked” once in a while to make it operate properly. But in today’s highly regulated environment, there’s always a question about whether or not such adjustments are allowed. For compendial methods in the pharmaceutical industry, the various pharmacopoeia give guidelines about method adjustment.
PEEK (poly-ether-ether-ketone) fittings and tubing are in widespread use for HPLC applications. The convenience of using your fingers instead of a wrench to tighten the fittings and the flexibility of the tubing make PEEK the material of choice for most HPLC methods when pressures of
As part of the HPLC Master Classes that I teach, I start each class by gathering questions or topics that the attendees would like to know more about. In a recent class in London, one of the topics was the causes of excessive retention – a method in which the retention times are longer than they used to be. There are a number of possible causes of increased retention in HPLC separations – let’s take a moment to highlight some of these. If we stop and think about it, there are just a few variables that control retention in HPLC. The primary ones are flow rate, temperature, mobile-phase composition, and column chemistry.
Ever have an HPLC method that specifies “ambient” as the column temperature? That’s a good sign of potential problems in the future for the method. We’ll get back to the interpretation of “ambient” in a bit, but first, let’s take a look at the influence of temperature on the separation.
In the last three instalments of HPLC Solutions, we’ve looked at the fundamental resolution equation
Rs = ¼ (N)0.5 (a-1) (k / [1+k]) (1)
In the last two instalments of HPLC Solutions, we’ve looked at the fundamental resolution equation
Rs = ¼ (N)0.5 (a-1) (k / [1+k]) (1)
Last week we began our study of the fundamental resolution equation
Rs = ¼ (N)0.5 (α-1) (k / [1+k]) (1)
Where N is the column plate number or efficiency, α is the selectivity, and k is the retention factor. We saw that, because N contributes to resolution as a square-root function, using N by itself is not a very powerful approach to make large increases in resolution, such as by twofold or greater.
In our last Back-to-Basics discussion (HPLC Solutions #39), we looked at the way we measure resolution, Rs, from a chromatogram using the following equation:
During method development in HPLC, one of the powerful variables that can be used to change selectivity is to switch from one solvent to another. For example, the B-solvent can be changed from acetonitrile (ACN) to methanol (MeOH) or tetrahydrofuran (THF). As a general rule, making such a solvent change will change the peak spacing, or selectivity. Hopefully this will improve the separation, but there is no guarantee of this.
If you use gradient elution for your HPLC separations, sooner or later you will run into problems with peaks that appear in the blank. Variously described as ghost peaks, background peaks or gradient interferences, these peaks can drive you crazy, especially if you are interested in measuring analytes that have small signal-to-noise ratios. Here we’re talking about peaks that show up in a no-injection, blank gradient. An example of an especially bad case of this is shown in the lower chromatogram of Figure 1. In this case, a 0.1% trifluoroacetic acid (TFA)/acetonitrile (ACN) gradient was run from 5% to 83% ACN over 13 min with a 5 min hold at the end. Although the none of the peaks are huge – the largest is just over 1 mAU in height – they were a problem for this method. These conditions were used for a stability-indicating assay that required reporting peaks of about 0.2 mAU – a real problem with this much background.
Q: I’m running an HPLC-UV method for routine analysis that has been in use for several years. Recently, I’ve been seeing more scatter in my calibration curves than I expect, and the quality control (QC) samples that I run with my method also are bouncing around more than normal. I suspect that there may be a problem with the pipette or the
pipetting process. Can you help me?
Q: I have a problem with the linearity of my HPLC method. The method involves derivatization of the analyte, followed by reversed-phase separation with UV detection. To my surprise, the calibration curve fits a quadratic form very nicely, and not a linear plot. Can you give me some advice on what I might be doing wrong?
Q: I’m hoping you might be able to shed some light on an issue we’ve had with unknown peaks in nearly every method we run in our lab. I’ve been working at µg/mL levels for a cleaning method for one product, while a colleague has been working at similar levels for a high potency product dissolution. The sample matrices are completely different, as are the LC methods; the only common denominator was the low UV detection (210–220 nm).
A reader recently sent me an email with this question: I run an LC-MS system and was recently tempted to try a recent published method using ion-pairing chromatography with 10 mM
tributylamine (TBA).After the trial, I converted back to the regular reversed-phase system and have now been plagued by the wretched 186 m/z peak for over a week. I have flushed the system thoroughly and replaced the peak tubing to no effect.
I doubt from this experience that I will be trying ion pairing again and would have hoped the publications would have included a warning regarding this effect. Can you offer any advice on how to purge this contaminant from the system?
Q: I have developed a method based on an article from literature for testing an analyte along with its impurities and degradants. This is a gradient method that utilizes a C18 column with 0.1% trifluoroacetic acid (TFA) in water (A) and 0.1% TFA in 80% acetonitrile/20% methanol (B). I tried other mobile phase additives to buffer the system (acetic acid, phosphoric acid, sodium phosphate) but using TFA resulted in the best peak shape and resolution. I’m not sure if that is the result of silanol suppression or ion pairing or both. However, I am concerned about (1) the stability of my column and (2) the stability of the TFA itself in the mobile phase. I’ve seen some baseline disruptions near the retention time of the analyte that ordinarily would be insignificant, but because I’m trying to show that the method can detect related substances down to 0.1% then it does become an issue with peaks near the lower limit of quantification, LLOQ. The issue with the baseline was not reproducible…it went away when I made a new batch of mobile phase. Do you have any recommendations for (1) extending the column life using TFA, and (2) minimizing baseline issues resulting from TFA degradation?
I recently had a reader ask me a question about what size the internal standard (IS) peak should be for a bioanalytical method (drugs in plasma). He was torn between advice that the IS should be about the same size as the smallest peak of interest, at the lower limit of quantification (LLOQ) and that it should be near the middle of the calibration curve. And if near the middle of the calibration curve, how is this determined with a method that has standards injected at concentrations of 1, 2, 5, 10, 20, 50, 100, 500, and 1000 ng/mL?
The following question was submitted by one of the readers of HPLC Solutions.
Q: I have a question about reporting the limits of my LC-MS/MS method. For this bioanalytical method (a compound extracted from eye tissue) the lowest concentration on the standard curve is 0.01 µM. The curve is made by spiking naïve eye tissue with the compound and extracting it before analysis. It is not a purchased compound, but is an endogenous compound which we’ve synthesized and use in our studies. All samples measured have higher concentrations than 0.01 µM, so it does not need to be lowered in order to bracket our samples properly. My question is, because I routinely detect this lowest standard and have a S/N of 33 at that level, can I call this my LLOQ? Does an LLOQ have to have a signal-to-noise of 10?
In previous instalments of this Back-to-Basics series, we have looked at the retention factor, k (HPLC Solutions, Issue #28 ), the selectivity, α (#31), and the column plate number, N (#32). Nice as these measurements are, they don’t tell us much about the quality of the separation. For example, you can have k in the ideal 2 < k < 10 region, or α = 1.2, or N = 15,000, but none of these values on their own make the separation any good. As a measure of separation quality, we need to determine the resolution, Rs, which is most commonly calculated as:
Rs = (tR2 – tR1) / ((0.5 * (w1 + w2)) (1)
Last issue (HPLC Solutions, #37), we examined how to calculate peak tailing. This issue, let’s take a closer look at some examples of peak tailing and see what they have to say in practical terms. For purposes of discussion, let’s consider the peaks in Figure 1, showing various degrees of tailing (asymmetry factor, As, used to calculate tailing).
So far in this Back to Basics series, all the peaks we’ve looked at are symmetric. The ideal peak in an HPLC chromatogram is Gaussian in shape, with an equal amount of distortion on the front and back edge of the peak. However, in the real world, this is rarely the case – most peaks tail. Because excessive peak tailing is an indication that something is wrong, it is a good idea to include a measure of peak tailing as part of the system suitability measurements.
Recently a reader sent an “Ask the Doctor” email to us asking what was meant when a column is referred to as end-capped, and what the function of the end-capping was. To understand end-capping, we need to step back and look at the bonded phase on the HPLC column. Reversed-phase HPLC columns usually comprise a silica particle with a stationary phase, such as a C18 hydrocarbon bonded onto the surface.
Previously we looked at the consequences of metal contamination of the silica used for packing HPLC columns. This week, we’ll consider another aspect of the silica – the nature of the silanol groups.
Today we often take for granted the quality of the silica in the HPLC column. This may be a reasonable assumption for the high-quality columns produced today, but it is not a guarantee, especially for columns that have been on the market for more than 10 years.
Earlier this spring, I was in London, teaching an HPLC troubleshooting class in conjunction with Separation Science . I love to teach, so the presentation of short courses is one of the highlights of my job...
In the past few articles we have been looking at some of the basic measurements that we can make to describe a chromatogram obtained from an HPLC method. So far, we’ve looked at the retention time, t R, the column dead-time, t 0, the retention factor, k , and selectivity, α. This week we’ll consider the last of the initial set of measurements, the column plate-number, N. This is also called the column efficiency, and is calculated as
N = 16 (t R / wb)2
where wb is the column width at baseline between tangents drawn to the sides of the peak, as in Figure 1.
An alternate way to calculate N is to use the peak width at half the peak height, w0.5:
N = 5.54 (t R / w0.5)2
We’ve been looking for the last few weeks at some of the calculations that are at the foundation of measuring the performance of an HPLC run. This week the topic is chromatographic selectivity. Selectivity is the ability of an HPLC method to separate two analytes from each other.Selectivity usually is abbreviated with the Greek letter α, and is calculated as: α = k2 / k1
Previously we have looked at the estimate of the retention factor, k , and the column dead-time, t 0. Here we’ll use some of the same numbers we used then to help diagnose problems with our HPLC separations.
In the previous article, we looked at the retention factor, k , and how to calculate or estimate it. In order to perform either of these processes, we need to know the column dead-time, t0. If we are using a UV detector and a “real” sample, usually there is an obvious disturbance in the baseline, as illustrated in Figure 1. If the sample is very clean, t0 might appear as a little zig-zag of the baseline [Figure 1(a)], but in most cases there is a significant “solvent” or “garbage” peak at the beginning of the chromatogram, as in Figure 1(b). As we’re only interested in a good estimate of t0 at this point, just pick a consistent place to measure it. The arrows in Figure 1(a) and 1(b) show where I’d pick to estimate t0 – just where the peak starts to rise above the baseline – this will be easy to measure consistently.
We’re going to take the next few weeks to visit the HPLC 1A class. We’ll be looking at some of the basic calculations used in HPLC, with an emphasis on their practical utility for evaluating separations, developing methods and isolating problems. The first in line is the retention factor, k, often called the capacity factor, k’. The retention factor is a measure of the distribution of the sample between the mobile phase and the stationary phase – somewhat analogous to a partition coefficient for a solute between two liquids.
I like chromatographic processes that can be simply described, wrapped in a neat package, and tied up with a pretty bow. There are a lot of cases where this is true, but not with column overload. Generally, we think of reversed-phase HPLC retention as non-polar solute molecules interacting with a non-polar stationary phase.
The recent discussion of various aspects of buffer use prompted one reader (D.B.) to enquire about how long one could use a buffer before it should be discarded. There are two aspects of buffer lifetime. One has to do with how long it is effective as a buffer and the second is related to other buffer properties – in particular, microbial growth in the buffer solution.
Shifting retention times within a sample batch can be a real source of headaches in the routine HPLC laboratory. Last week we talked about the kind of retention drift experienced by some HPLC methods for the first few injections. This often is associated with slow equilibration of a mobile phase or sample component with the column, but usually disappears after the first three or four injections. We saw that often a large-concentration loading injection would speed this process and that ignoring the first injection or two was the simplest way to deal with this problem.
How many of you have experienced this? You think you’ve fully equilibrated the HPLC column by running mobile phase through it for 30 min, but for the first 3 or 4 injections, the retention times drift a little, finally settling down after half a dozen injections are made. In some cases, the problem occurs only with a new column, whereas in other methods, each time the method is started, the same pattern of drift is observed.
My short answer: Yes!... and No!... and Maybe. A guard column is one of those HPLC accessories that is a bit of a mixed grill – it can be a good thing or a not-so-good thing. Let’s look at some of the different aspects of guard column use to see if they make sense for you.
In the previous instalment, we looked at the processes of sample filtration or centrifugation as a means of reducing or eliminating particulate matter from the injected sample. As I mentioned, I prefer centrifugation, because it is easy, inexpensive and effective. However, it is not perfect. This means that it is possible to get some particles injected along with the sample. And the sample is not the only source of particulate matter that may enter the HPLC column. Worn parts,especially the pump piston seals and the injection valve rotor seals, also can contribute to the particulate load of the system. Although it is pretty easy to minimize pump seal degradation by regular replacement on a 6- or 12-month cycle, injection valve seals seem to last forever – they can last 100,000–500,000 cycles or more. But when they fail, they can shed particles that can block your column.
It is easy to take knowledge for granted, especially when you’ve been involved in a field for a long time, as I have with HPLC. One of those areas is about how to protect the column. In the training courses we teach, Tom Jupille and I jokingly instruct the attendees that the best way to make a column last forever is to leave it in the box. But this isn’t very practical, so for the present discussion, I’m going to concentrate on how to prevent particulate matter from getting to the column.
A few weeks ago, we spent several issues of Solutions (12-14) talking about various aspects of buffers. We saw that some buffers were better than others and that there were certain buffer-preparation practices that should be avoided. This prompted one reader (I.M.) to remind me of another wise practice relating to buffers – to anticipate the use of an LC-UV method with LC-MS detection. This also reminded me of my years as a Boy Scout and scout leader – and that the Boy Scout Motto is “Be Prepared.” This is a good motto for chromatographers, too.
Sometimes I don’t know whether to be amused or dismayed when I am presented with an HPLC method. Whether it is a method question from one of the readers or a paper to review for the 'Journal of Chromatography', it is a bit surprising to me how often methods undergo a process that I refer to as “genetic drift.” This kind of method results from constant tweaking of existing methods to perform new tasks. Let’s look at some examples of such methods.
HPLC Solutions #18: My Favourite Things, #1: “Introduction to Modern Liquid Chromatography” 3rd Edition
My first “real” job was working for Technicon Instruments Corp. in Tarrytown, New York, USA. My boss was Lloyd Snyder, who was to become a close friend and who has been my business partner for almost 26 years. Little did I know what my future had in store when he gave me an assignment of reviewing the draft of Introduction to 'Modern Liquid Chromatography', which he and Jack Kirkland had written for the 2nd edition (published in 1979). Without a doubt I learned more about chromatography while reviewing that book for typos than I have from any book before or since. When I say that the cover is falling off of my copy, I really mean it – there is a tear down the cloth of the spine from being pulled off the shelf so many times. It still has the original dust jacket, but it has been taped so many times there is more tape than jacket in most places. It still sits proudly on my bookshelf next to its older brother, the 1st edition, published in 1974.
Save 50% on the cost of your LC-MS! Now that I have your attention, perhaps I shouldn’t make such a grand claim – maybe only 40%. Of course I’m talking about operating costs, not the purchase price, but if it means that you can do the work of two LC-MS units by making more efficient use of a single LC-MS, isn’t this the same colour of money?
Q: Is it OK to back-flush a column? I’m using a C18 column for a routine analysis and the pressure has risen. I think that if I reverse the column I can unplug it. Is this OK or will I damage the column.
Q: I have observed peak splitting in my chromatograms when I inject the sample in tetrahydrofuran (THF) as the injection solvent, but I don’t see any problems when acetonitrile (ACN) is used. I am using a C18 column with a mobile phase of ACN/buffer and an injection volume of 50 µL. Do you have any idea what could be going wrong?
Q: What buffer should I use when I want to optimize the pH of the mobile phase? I normally use phosphate buffer, but I heard that phosphate may not always work. How can this be? I thought that phosphate was a great buffer for reversed-phase HPLC. Can you clarify this?
When a buffer is used properly, the mobile-phase pH will stay constant even when assaulted by the sample or injection solvent. For this reason, buffered mobile phases are strongly recommended for most reversed-phase HPLC applications.
Q: I use trifluoroacetic acid (TFA) to adjust the aqueous portion of my HPLC mobile phase to pH 2.5. Even with careful pH adjustment, I see changes in retention when I prepare each new batch of mobile phase, such as shown in Figure 1. Retention seems to be consistent within a batch, but batch-to-batch variability concerns me. Can you shed any light on this problem?
Q: I know that washing an ion-pairing reagent from the column can be difficult. What’s the best way to do this?
Q: What is the relationship between pore size and particle size for an HPLC packing material?
PEEK (poly ether ether ketone) tubing has become a standard item in the operation of many HPLC systems. It is convenient, inexpensive, and easy to identify the internal diameter. However, some care needs to be taken to avoid problems associated with PEEK tubing
Q: I accidently ran out of solvent in an overnight run on my HPLC system. I had not activated the low-pressure limit on the pump, so the pump just pumped air for half the night. Will this ruin my column? If not, how can I get all the air out?
Q: I know that retention varies according to the ionization of a compound. If the compound is ionized, the retention generally is shorter than if it is not ionized. Does this mean that I should expect two peaks if the compound is near the pKa, where it is half-ionized?
Phase dewetting in HPLC is the same process we used to call phase collapse. Although often drawn to look like a tennis ball with stationary phase with the C18 groups as fuzz on the surface, the particles used to pack a C18 column are totally porous, resembling a rigid sponge.
There are two procedures that are used to make sure that an HPLC equipped with a mass spectral detector (MS) is reporting proper mass information and that the system is properly set to detect the analyte(s) of interest.
The signal-to-noise ratio (S/N) is an important variable that may influence the performance of your method.
I often get asked which HPLC system or manufacturer is the best. In days of yore, there was quite a bit of difference in quality and reliability between instruments sold by different manufacturers, but that is no longer true.
Q: Which is better, high-pressure mixing or low-pressure mixing?
Q: I have a validated stability-indicating assay that uses one manufacturer’s C18 column. I know of another column manufacturer where I can get an equivalent C18 column for half the price of the column stated in the method. Will changing columns invalidate the method?