HPLC PEAK Fronting and Tailing, Common Reasons For It

All users of HPLC need to know and be familiar with the correct terms used to describe non-Gaussian shaped peaks. Two of the most common undesirable peak shapes, peaks that show "Fronting" and peaks that show "Tailing" indicate problems with the HPLC method.  A quick refresher on why you may observe an HPLC peak front or tail on the chromatogram follows. 

Peak FRONTING: First, let us define what peak fronting looks like. The leading edge (front) of the peak is vertical, straight up and non-Gaussian in shape. This sharp increase in signal is easy to spot. 

Common Reasons for Peak FRONTING:

• Poor sample/peak capacity. In other words, too low a K prime (not enough retention on the HPLC column) resulting in no chromatography taking place. To solve this problem you must develop a proper HPLC method which first retains the compound(s) of interest, holds them long enough to obtain an acceptable K prime and resolve them away from other peaks, then elutes them off the column.

• Injection Solution Too Strong:Your sample(s) should be dissolved in the mobile phase and not in a solution that is "stronger" in elution strength than the mobile phase. Example: If you method is 100% aqueous, do not inject the sample in a solution with organic solvent. Follow fundamental good chromatography guidelines.

• Column Fouling / Overloading of sample. When the HPLC column is overloaded with sample, the peak shape will show fronting. Decrease the injection volume and/or concentration, as appropriate, in 10x graduations until the peak shape is normal.

• Saturation of the Detector: Just as with overloading the column the peak shape may change, overloading the detector's measuring range may also result in saturation of the signal and loss of accuracy. Decrease the injection volume and/or concentration, as appropriate, in 10x graduations until the peak shape is normal and back on-scale.

Peak TAILING: First, let us define what peak tailing looks like. The trailing edge (tail) of the peak slowly drops off towards the baseline and  is non-Gaussian in shape. For those with GC experience it appears similar to a peak that "bleeds" and continues to interact with the column for an extended period of time.

Common Reasons for Peak TAILING:

• Flow path Diffusion (from extra-delay volume). Poorly swaged fittings/connectors, a column with a void, incorrectly sized capillary connection lines may all contribute to peak tailing. Optimize the flow path, column and connections.

• pH dependence for ionizable compounds. If the sample is easily ionized and the difference between the pka of the sample and the mobile phase is less than 2 pH unit, tailing may result. Being sure to work within a safe pH range for your column, increase or decrease the mobile phase pH to be > 2 pH units away from the sample's pka to reduce tailing.

• Type 'A' silica or heavy metal contamination of the support. Many older style column supports did not use ultra-pure, heavy metal free packing material. These material often interacted with the sample on the column resulting in changes in retention, The use of more modern type 'B' or 'C' packings has eliminated many of these problems.

• Residual silanol groups present on support. As with the earlier type 'A' supports, non fully end-capped supports with residual silanol groups often resulted in secondary, extended retention effects. Use of more modern, fully end-capped, ultra-high purity packing materials (and/or mobile phases which better address these residual groups) often allow Gaussian peak shapes without the need for many additives.

• Column Fouling / Overloading of sample. When a column is not washed of all retained material after each analysis, it may build up over time and change the surface chemistry of the support. This may lead to changes in retention, especially delays in both binding and elution. Wash, regenerate or replace the column to solve.

You may also be interested in reading a related article; "Two Common HPLC Problems and their Causes (Sudden changes to either the HPLC Backpressure or Peak Shape)".

Standard Addition HPLC Calibration Method (Spiking)

The standard additions calibration method is used to determine the concentration of an analyte which is in a sample matrix such as is commonly found when working with clinical, biological or food samples. To rule out interference from other components in the sample matrix, the sample is “spiked” and the detector response is monitored for any change which is not due to the change in sample concentration. The fortified standards created by spiking the original samples is useful when no version of the sample matrix is available without the analyte for use as a true blank. 

Common “Spiking” Standard Addition Procedure Example:

Divide up your sample evenly into FIVE volumetric flasks, representing five different final concentrations of spiked samples. Into four of the flasks add the same volume of increasing linear levels of the analyte to create four different calibration levels (e.g.  spiked amounts such as 25, 50, 75 and 100 units). Into the fifth flask add the same volume of diluent, without any spiked component, to serve as the ‘0’ or blank equivalent std. Next, extract all samples, inject and analyze the data per the usual method. A calibration curve of this new data set should show a linear relationship. If so, and your method is validated, you may be able to use just one spiked sample along with your regular samples to show that the reported concentration of your sample is not effected by the matrix and save yourself a lot of work (because you would need to prepare these standards and run a calibration curve each time).

Carry-Over (Carryover) Contamination in HPLC and LC-MS Systems

"Carry-over" is a term used to describe a type of sample contamination which causes sample peaks to re-appear in later runs which do not actually contain the sample (e.g. blank runs). The contamination can last for several sequential runs, often decreasing in amount after each injection (which is a key observation when troubleshooting). When proper instrument training has been provided, modern HPLC system designs make carryover extremely rare, but when it does appear, the contamination can be due to: (1) A lack of HPLC maintenance; (2) Overloading samples which foul the column; (3) Poor Wash Vial Usage and/or Sample Vial Selection; (4) Inadequate operator training in how to set-up and use the chromatography system. *Note: Proper operator training greatly reduces the chances of contamination and is the most overlooked reason for the problem.

The Types of HPLC Carry-Over Contamination; Why They Occur and How To Reduce Them:

(1) A Lack of HPLC Maintenance: Most auto-injector valves rely on a rotary seal to move the sample from the needle loop to the flow path of the system. The components within these valves wear out and should be inspected at least every 6 months and replaced when needed. Also, always check the needle seat and needle for signs of wear or leaking. Note: Look for signs of leaks by the injector. Leaks always indicate a problem and should be fixed immediately. Don't run samples when you have leaks. Your method and data obtained will be invalid. Any worn parts should be replaced and the system performance tested. One of the most common causes of carry-over is due to a worn sample injector valve rotary seal. A worn seal can allow sample to be retained in the worn grooves, in-between injections, resulting in sample peaks appearing in subsequent runs. Additionally, buffer salts can lodge between the seals causing leaks or carryover. Routine HPLC service and, if applicable, flushing of all buffers/salts every day can eliminate these issues.

(2) Column Fouling / Overloading of Sample: If you inject too high a concentration of sample and overload your column with material, then it is possible that your column will continue to bleed sample long after the analysis is over. This also happens when the sample has a high affinity for the support you have chosen too. Failure to regularly flush and clean your HPLC column on a regular basis can also result in a similar problem as retained material is released from the column over time. Avoid this problem by performing a loading study to determine how much material can be effectively loaded on to the column. Next, create a wash method which utilizes a stronger solvent than your method (often utilizing a gradient) which will wash away any strongly retained material in between runs. This is critical if you are running an isocratic method as material will be retained on the column and must be washed off at frequent intervals using a stronger wash solution. *When using only isocratic methods, people often do not initially observe carry-over problems (because the sample just sticks to the column and accumulates over time). When the solvent strength is changed or the method is revised to a gradient, then the problems start... Avoid the problem by selecting the right column (which retains, then elutes ALL of the sample), not overloading the column (do a loading study) and washing the column down with a stronger solution that fully dissolves (not precipitates out) any remaining material off the column after each run.

(3) Wash Vial Usage and/or Sample Vial Selection: If you are using a modern high-pressure, "Flow-Through" design autoinjector (e.g. Agilent 1100, 1200, 1260, 1290), then carryover is rarely an issue as these modern injectors use a high pressure pump to aspirate and inject the samples directly into the flow path, reducing the need for any wash stage. The entire HPLC's injection flow path is continuously washed with mobile phase during the analysis run. This dramatically reduces the chances of any sample re-appearing in later runs. The need for a separate wash vial is nearly eliminated in this way as the needle, needle seat, loop, injector pump and valve are all flushed clean during each method. Many older auto-injector designs utilize either a low pressure injector (glass syringe) or injector pump which is not part of the main flow path. These injectors benefit from a separate wash vial as they are not continuously cleaned. Effective cleaning requires that a wash vial be employed in these cases. It should be filled with mobile phase or a solution which will dissolve any remaining material which might still be in the system.

When sticky sample solutions are used, separate Wash Vials can be used to reduce contamination with either older or newer injector designs . Sometimes these sticky samples can adhere to the outside of the needle while it is being withdrawn from a vial which has a septa which has been punctured many times. High puncture rates tend to open up the hole resulting in a lack of "wiping' of the needle surface upon withdrawal. *For vials that are punctured many times, it is critical to replace the septa OR use septa materials which seal for a long enough time frame to reduce this effect. Septa needle wiping eliminates some of this contamination. Two types of contamination can occur from this problem. (a) When the needle is dipped into a vial (same or different one) which also has a large septa opening, it can carry some of the sample with it and deposit it into the new vial (or onto the septa of the vial). (b) The contamination can also run down the needle itself and drip onto the needle seat at the time of injection resulting in contamination of the seat or sample.

One of the easiest solutions to reduce external needle contamination involves incorporating a wash vial which contains a solution which is optimized to quickly dissolve the sample into solution. This sounds simple, but many chromatographer's choose wash solutions which do not enhance the cleaning aspect of the needle at all. For example: Mobile phase, which is normally ideal, but does not work in some cases. Samples such as peptides, proteins, fats, oils and/or lipids can be troublesome as their solubility can be at odds with the mobile phase chosen. For the wash vial to be effective, it must quickly dissolve the material. The needle can be first "dunked" (dipped) into one vial containing the solution and withdrawn, followed by an aspiration and wash in a second wash vial. If needed, you take this cleaning one step further and use additional aspiration steps to serially dilute any remaining material. These wash vials must be changed frequently (easily done by having several wash vial positions programmed in the system). Additionally, the caps should be left OFF the wash vials to reduce pickup contamination each time they are used (this step is critical).

Lastly, if you are analyzing sticky materials which are known to interact with metals found in chromatography systems, consider using a system which incorporates bio-compatible materials such as titanium, tantalum and/or polymers such as PEAK. You can also utilize plastic sample vials (e.g. PP) or plastic vial inserts too.

(4) Inadequate Operator Training: Good chromatography requires a complete understanding of the hardware used and the fundamentals of HPLC. You must be able to troubleshoot the complete flow path of the system and understand the concepts of chromatography as used in method development. This is not a technique best learned by trial and error, but rather through mentoring using logical steps. Depending on your skill set, troubleshooting a "carry-over" problem in an HPLC system can take minutes to months to diagnose and solve. We learn these skills through hands-on experience and training. Reading many of the better books and articles on the subject matter helps too. Get as much practical hands-on training as you can. Ask your supervisor or manager(s) to invest in you by purchasing professional training for you in this field so you can learn on your own systems. You will learn far faster this way and spend less time troubleshooting problems and more time running samples, accurately in less overall time. Training also costs just a fraction of what the instrumentation and your salary are. If you have acquired the fundamental skills, a skilled teacher can impart about one years worth of practical knowledge to you in as little as one week of training.

Summary: The two most common reasons for sample carry-over contamination in an HPLC or LC/MS system are due to: lack of operator training and/or lack of system maintenance (most commonly manifested as a worn injector rotor seal).

 Note: This article specifically addresses carry-over contamination as it relates to the most commonly used HPLC, UHPLC and LC-MS autoinjectors (or autosampler modules).

You may wish to read a related article on "Troubleshooting HPLC Injectors (Manual and Automated)" found at this link: http://hplctips.blogspot.com/2013/06/troubleshooting-hplc-injectors-manual.html

Syringe Filter Selection for HPLC or LC/MS samples

This article will address the use of disposable female, Luer-compatible, syringe filters without built-in pre-filters for the filtration of individual samples into vials for HPLC or LC/MS analysis. - Note: 96 or 384 multi-well filtering plates provide for a better solution when large quantities of samples need to be filtered. Note: The presented filter membrane material selection criteria also applies to mobile phase filtration too.

The choice of syringe filter depends on the: filter size (volume) of your sample, the chemical compatibility of the housing and membrane and desired pore size. Selection of the wrong filter size can result in too much sample holdup volume (loss of sample on filter) or overloading of the filter (allowing unfiltered material to pass through). If a membrane or housing is chosen which is not chemically compatible with your solution, then contamination of the sample or rupture of the assembly can result. Choosing a filter with too large a pore size can result in material passing through it which could clog or contaminate the solution (i.e. plug an HPLC system or result in a loss of sterility of a solution). Protein binding affinity is another characteristic of filter membranes and if you are filtering samples of biological interest, then you will also want to consider this specification in your selection criteria too (though it will not be discussed in this article).

Syringe Filter Size:

Filters are available in a variety of sizes which are generally in a disc shape and described by their diameter. Common sizes available for chromatography samples include: 3 mm, 4 mm, 13 mm and 25 mm (~25 - 30 mm) diameter discs. The larger the diameter of the disc, the larger the sample capacity, cross sectional surface area and potential hold-up volume of the sample on the filter. 

Hold-up volume is important because some of the sample will be retained inside the membrane and/or filter housing. If too large a filter is selected, samples with small volumes could be lost entirely in the hold-up volume on the membrane. Smaller filters have lower hold-up volumes. To extract as much sample as possible, be sure and use a post-filtration air purge to reduce the total hold-up volume.

If the volume of the sample you wish to filter is under 1 ml, then a 3 mm filter may provide the lowest hold-up volume and require the smallest amount of solution. To filter samples between 1 ml and 10 ml, the 13 mm diameter filter provides a good balance between hold-up volume and large filter surface area. Larger sample volumes from 5 ml to 50 ml are often filtered through the more common 25 mm diameter filters (~4 times the filtration area as a 13 mm disc).

Chemical Compatibility:

Membrane Material: This is where you really must consult the manufacturer’s own documentation for the most compatible filter membrane for both your sample and the solution that will flow through the filter. To simplify the selection criteria, we can make some generalizations about some of the different types available:

Cellulose Acetate (CA): Use with aqueous solutions and a few hydrocarbons only. Low protein binding so good for many biological samples. Not compatible with ACN or DMSO. Can be autoclaved.

Nylon: Great general purpose material and compatible with many HPLC solvents (including THF, alcohols, ACN), but not strong acids. Nylon has a high affinity to bind proteins. Can be autoclaved.

Polysulfone / Polyethersulfone Variants (PS / PES): Commonly used with tissue culture and ion chromatography samples. Stable with many strong bases and alcohols, but few HPLC solvents (as it is hydrophilic). Low backpressure and low protein binding. Not compatible with ACN. Can be autoclaved.

Polypropylene (PP): General purpose hydrophilic material with resistance to most acids, bases, DMF, DMSO and alcohols. Not recommended for use with hydrocarbons, esters or solvents such as ACN. Can be autoclaved.

Polyvinylidene difluoride (PVDF): Hydrophilic material with broad compatibility. Often a good choice for use with alcohols, hydrocarbons, biomolecules, ether and ACN. Low protein binding. Can be autoclaved.

Polytetrafluoroethylene (PTFE): Reported in most brochures to be chemically resistant to almost all solvents, strong acids and bases. Hydrophobic membrane should be pre-wetted when used with aqueous solutions. Low protein binding and very strong. Can be autoclaved.

Most chromatography grade syringe filters are constructed of either HDPE or PP. These materials are compatible with a wide range of HPLC solvents and both offer low levels of extractables. HDPE has been reported to be more chemically compatible with aqueous basic solutions of NH4OH than PP.

Pore Size:

This will depend on your application and a number of different pore sizes are commonly available from vendors (1 micron, 0.8, 0.45 and 0.22 micron are the most common): 

For example, is sterilization of the fluid the goal? If so, a 0.22 micron filter is generally accepted as the best choice.  


For most chromatography or LC-MS applications either a 0.45 or 0.22 micron filters are preferred.

Summary:

• Please refer to the various manufacturers data sheets to select an appropriate syringe filter with: (1) a low hold-up volume; (2) large enough size for the volume of sample; (3) which is chemically compatible with the solution and material you are going to inject through it and (4) lowest protein binding affinity (if applicable).
   
• To reduce the hold-up volume, use a post-filtration air purge to empty the filter.
   
• Minimize contamination from extractables (in the plastic) by pre-rinsing the filter membrane with some of the clean solution. This can reduce the amount of detectable extractables in your sample. PTFE based membranes have some of the lowest extractable levels so consider their use if this is an issue.
   
• If analyte binding is a concern, select one of the membranes which has the lowest binding affinity such as PVDF or PTFE.