Dept. Analytical Research and Development, Pfizer Worldwide Research and Development, Sandwich, Kent, UK,

Dept. Forensic Science & Drug Monitoring, King’s College London, London, UK, 

Present address: Dept. of Chemistry, University of Southampton, Southampton, UK.

This article presents a method for comparing the levels of baseline interference arising from common laboratory mobile phase contamination sources and assesses different approaches for removing dissolved contaminants to generate interference-free chromatogram baselines. The authors demonstrate that recirculating mobile phase through a semi-preparative scale column using a reagent delivery pump has advantages over previously published mobile phase decontamination methods. 

	Increasing regulatory scrutiny combined with improved modern liquid chromatographs used in pharmaceutical purity analysis has seen a trend towards increased detection sensitivity. It is now commonplace in pharmaceutical analysis to measure impurities and degradants at sub-nanogram concentrations from sample column loads of 200 nanograms or less when using UV detection. When performing such high-sensitivity measurements, complications with background environmental contamination of the system and mobile phase are often encountered. Commonly referred to as ghost peaks but more properly denoted as contaminant peaks, extraneous signals arising from UV absorbing contaminants in the liquid chromatography (LC) system can interfere with measurements of low-level solutes. Unless special measures are taken the interference from contaminant peaks can lead to inaccurate measurements of low-level impurities. 

	Modern chromatographs with low dead volumes and high sensitivity UV detectors are commonly used in pharmaceutical analysis (1,2,3) with gradient elution to separate diverse polarity solute mixtures of active pharmaceutical ingredients (APIs) and impurities within reasonable run times (4). UV detection is still commonly preferred to mass spectrometry (MS) because of reduced cost and ease of methods transfer. Laboratories are often required to quantify solutes at sub-nanogram concentrations (5) when measuring low levels of potentially genotoxic impurities (6) and testing the effectiveness of cleaning methods (7). According to current industry guidelines (8), methods for the purity analysis of pharmaceuticals should be capable of quantifying impurities above a 0.03% threshold for drugs with a maximum dose of more than 2 g/day. For doses of less than 2 g/day the threshold is relaxed to 0.05% of API concentration. In early development stages reference standards are often unavailable for all solutes of interest and a peak area normalization approach is commonly used to monitor and control impurity levels. When working at such extremes of sensitivity in industrial environments it can be difficult to avoid interference from environmental contamination unless special measures are taken.

	In modern industrial settings detection of unwanted extraneous contaminant peaks in the chromatogram baseline can interfere with measurements (9,10) and require additional time and effort to identify the cause and implement counter-measures. These can have a significant impact on the transferability of analytical methods because of differences in the purity of reagents and handling practices at each site - often the causes remain unresolved (11,12). Extraneous peaks in the chromatogram that are not directly related to the injected sample of interest have been referred to as system peaks (13), vacancy peaks (14), artefact peaks (15), and induced peaks (16). However, the most commonly used term is “ghost peaks” (17,18), and because they usually arise from some kind of system contamination it is more correct to refer to them as “contaminant peaks” (CP).

	Since the advent of gradient liquid chromatography (LC) baseline interferences arising from mobile phase contamination has been a problem and many publications have been devoted to this subject (19,20,21,22,23,24). Gradient elution exacerbates contamination problems because trace contaminants become enriched on the column and then elute once the elutropic strength becomes sufficiently high. There are lots of possible sources of contamination in the modern LC laboratory and trace contamination can be very difficult to avoid. Contamination of water with airborne particles, organics, colloids, and ions (11,25), impure additives (19-24,26), dissolved gases, and organic solvent impurities (11,27,28) have all been reported, as has accidental contamination from pH meters and laboratory glassware (19-24). 

	It has been recommended that LC mobile phases should be freshly prepared and micro-filtered before use (29) and that LC solvent lines and glassware be regularly rinsed to avoid build-up of biofilms and detergents (30) and because of the propensity for microbial growth in buffers. Plasticizers such as phthalates are prevalent in the environment where indoor air levels have been detected above 10 µg (31). Phthalates can be detected on most surfaces and are readily extracted by organic solvents (32). In a laboratory setting plasticizers were detected in laboratory water and leaching from consumables (33). Adsorption of plasticizers from the air can compromise the long-term stability of reagents (34) and the concentrations of these contaminants can be increased by exposure to laboratory equipment and manual handling steps (35).

	To remove contaminant interference in LC various methods for contaminant extraction from the mobile phase have been published. Solid-phase extraction (SPE) using hydrophobic sorbents has been applied with some success (36,37,38). However SPE under vacuum is prone to flow-channelling and breakthrough of more polar impurities, particularly with membrane disks, which have limited retention capacity as a result of short bed length (39). Variability in contaminant removal effectiveness has also been reported (40,41). 

	Mobile phase decontamination using in-line guard columns helps to remove micro-particulates and prolongs the lifetime of analytical columns. However, because they remain in-line to the analytical column, they are not a reliable way of removing contaminant peaks. The use of high performance liquid chromatography (HPLC) columns as impurity traps (installed between pump and mixer) has been shown to be effective at removing dissolved trace contaminants from the aqueous phase, resulting in reduced levels of baseline interference, but the use of short trap columns limits the effective lifetime of the trap before contaminant breakthrough occurs (42). This is particularly true for more polar contaminants, which can break through the trap columns and refocus on longer analytical columns to cause interference.

	The use of longer, larger capacity trap columns can be limited by the undesired contribution to extra system volume if applied directly to the analytical LC pump. To do this efficiently the trap column needs to be of sufficient scale to retain contaminants from a large number of gradient cycles and operate throughout a run sequence. Using a reagent delivery pump it is possible to perform mobile phase decontamination outside of the flow-path of the analytical LC system. In this novel configuration the trap column does not contribute to system back-pressure or dwell volume and can be applied with any type of gradient LC pump. On-line continuous clean-up of mobile phase using longer bed length and larger capacity trap columns protects against subsequent airborne contamination without the need to flush the trap between injections. 

	This article presents a method for comparing the levels of a LC-UV chromatogram baseline interference arising from mobile phases exposed to different contaminants and decontamination methods in an attempt to highlight the highest risk sources of mobile-phase contaminant peak generation and test the effectiveness of various approaches for decontamination of the mobile phase. 

The (95% v/v) aqueous mobile phases were prepared using ammonium acetate (HPLC grade, Thermo Fisher Scientific) to a concentration of 10 mM with addition of acetonitrile (5% v/v, Sigma-Aldrich). Purified water was obtained from a Milli-Q Gradient A10 Purification system (Merck Millipore). Mobile phase solutions were stored at ambient laboratory conditions during the evaluation. Larger batches were stored in 10 L polypropylene bottles and smaller sub-lots used in systematic tests were stored in 1 L capacity clear glass Duran bottles that were washed thoroughly with organic solvent (acetonitrile) and allowed to dry (inverted) before use. 

 To determine the sources of potential contamination, mobile phases were deliberately contaminated by objects commonly found in the laboratory: a plastic weighing boat, dishwasher detergent, Tygon tubing, Parafilm, contaminated pipette, and two different types of nitrile glove; the thin single-use disposable type often used for general laboratory tasks and the thicker reusable type. Dust particles were collected from our own laboratory windowsill. A 5-L batch of ammonium acetate buffer (native pH measured at 6.8) containing 5% acetonitrile was prepared and 500-mL lots were subdivided into clean glass bottles, which were rinsed out before use with high-quality acetonitrile. Each bottle was deliberately contaminated by immersing solid items (for example, gloves, Parafilm) in the mobile phase for 20 s. A small pinch of dust (c.a. 0.5 mg) was sprinkled onto the surface of one bottle for comparison. Each contaminated mobile phase lot was used to generate a baseline following enrichment of 60 mL of the mobile phase onto the column before running an acetonitrile gradient program to elute the contaminants. To test the efficacy of the clean-up techniques under worst-case conditions, a 5-L stock solution was prepared and then sub-aliquotted into 500-mL lots, each of which was deliberately contaminated with the common laboratory products mentioned previously. The mobile phase was tested before and after each of the different clean-up techniques and compared to an unadulterated batch of the stock solution.

To test the purity levels of water and acetonitrile a number of different grades and sources were obtained to test any differences between suppliers and grades. Water was also obtained from a Milli-Q Gradient A10 purification system (Merck Millipore). 

 Emporestyrenedivinylbenzene (SDB-XC) 47-mm SPE filter disks were obtained from Supelco. Oasis Hydrophilic Lipophilic Balance (HLB) (6 g) packed particle SPE cartridges were obtained from Waters, and Envi-Carb 12-mL (1 g) and 3-mL (0.25 g) porous-graphitic carbon (PGC) SPE cartridges from Sigma Aldrich. For the off-line SPE techniques, a vacuum manifold (Waters) was used. The adulterated mobile phase (500 mL) was passed through each SPE cartridge following pre-conditioning with methanol (6 mL) and purified water (6 mL) and collected in a glass bottle pre-rinsed with acetonitrile. For the filter disks, a glass vacuum filtration apparatus (Millipore) pre-cleaned with acetonitrile was used to perform the filtration. Each SPE device was pre-washed with acetonitrile (10 mL) and purified water (10 mL) before passing 500 mL of mobile phase through under slight vacuum. Stir-bar sorptive extraction (SBSE) stir-bars (4 cm, coated with polydimethylsiloxane (PDMS) were donated by RiChrom. The extraction was performed by stirring 500 mL of aqueous mobile phase at a medium speed (c.a. 120 rpm) for 60 min before sampling. All treatments were prepared in singlicate.

In-Line and On-Line Extraction Techniques:

 A Turboflow Cyclone-MCX column (4.6 × 100 mm, 50-µm) containing a styrene divinylbenzene (SDB) copolymer and a Turboflow Silica-C18 column (4.6 × 100 mm, 50-µm) were donated by Cohesive Technologies. Ghost-trap DS micro-columns (4.0 × 20 mm) were purchased from Shimadzu. Luna A C18 (2) semi-preparative column (150 × 10 mm, 5-µm and 150 × 21.2 mm, 5-µm) were obtained from Phenomenex. The trap columns were evaluated by placing them (a) in-line between pump-head and mixer (for turbulent flow chromatography (TFC) columns only) and (b) connecting the C18 columns on-line by pumping into the mobile phase inlet line using an additional model 422 isocratic pump (2 mL/min, Kontron) via a tee-piece with an overflow line back to the reservoir. 

	Oxycodone (OC, Sigma Aldrich) standard testing was performed using Agilent 1100 series HPLC systems comprising of an on-line degasser, low pressure mixing quaternary pump with proportioning valve, auto-sampler, and diode array UV detector with Empower software control (Waters Inc.). UV detection wavelength was 210 nm. Standards were prepared at concentrations of 200 µg/mL and 100 ng/mL and 10 µL injections were made onto a 4.6 × 150 mm, 3.5-µm XBridge C18 column (Waters Inc.) at a flow rate of 1 mL/min. Mobile phase was ammonium hydroxide (Fluka LC-MS grade 1 mL phials, Sigma Aldrich, diluted to a final concentration of 0.1% v/v with purified water) and acetonitrile. A gradient elution programme was performed by changing the acetonitrile composition from 5% to 64% over 29 min following a 2.5 min hold at 5% acetonitrile with a flow-rate of 1 mL/min. Mobile phase composition was maintained at 64% acetonitrile until 34.5 min, after which the column was re-equilibrated at 5% acetonitrile for 5 min. Column temperature was maintained at 32 °C, which was previously determined as the optimum for the separation of OC and its impurities during method development activities.

 HPLC separations investigating the sources of mobile phase contamination and the effectiveness of removal methods were performed using two different Agilent 1200 series HPLC systems comprising of an on-line degasser, binary pump with high-pressure static mixer, autosampler, and diode array UV detector with Chemstation software control (Agilent Technologies Inc.). UV detection wavelengths of 210, 240, and 254 nm were used for detection. Chromolith C18 monolithic HPLC columns (10 × 4.6 mm, obtained from Merck) were used for the trapping, elution, and resolution of enriched contaminants. To study the sources of contamination, zero volume injections were performed following equilibration of the column for 20 min at a flow-rate of 3 mL/min and organic composition of 5% acetonitrile. Following equilibration a gradient ramp programme of 5-95% acetonitrile over 10 min was used to elute the enriched contaminants from the column. For the evaluation of clean-up techniques, the equilibration time was reduced to 5 minutes prior to running a gradient of 5-95% acetonitrile over 10 min. Duplicate measurements were made for each tested lot of mobile phase. The first measurement was discarded and the second replicate was used for the comparison. To avoid cross-contamination systems were washed by flushing lines with 80:20 (v/v) acetonitrile:water before use. Gradient-grade (≥99.9%) acetonitrile (Sigma Aldrich) was used throughout the clean-up and contamination experiments. Trap columns of all sizes were prepared by flushing with MS-grade acetonitrile for 15 min followed by water (obtained from the  MilliQ system) containing 5% acetonitrile for 15 min at 1 mL/min (micro- and analytical-scale traps) and 10 mL/min (semi-prep and preparative column traps).

	To be able to measure and compare the level of contamination arising from different mobile phase treatments, a method was developed to equilibrate a column with 60 mL mobile phase under highly aqueous conditions before eluting contaminants using a gradient elution cycle. Baselines were monitored at a range of wavelengths typically used in pharmaceutical analysis by LC-UV. Comparison of different contamination sources by exposing mobile phase to different contaminants confirmed that common laboratory equipment could have a significant impact on the formation of contaminant peaks. Example chromatograms from these studies are provided in Figure 1. All tests showed some degree of contamination would accumulate if more than one contamination source was exposed to the mobile phase during preparation.

	Disposable nitrile gloves are frequently used in many LC laboratories. Though it would not be considered normal practice to immerse a whole glove into a mobile phase container, it is common to handle mobile phase transfer lines, filter sinkers, bottle necks, and caps whilst wearing these gloves, and the high level of mobile phase contamination observed from this source indicate that they are a potential risk for generation of unwanted contaminant peaks. Dust contamination of the mobile phase also generated a significant increase in the contaminant peak profile. Dust is always present in laboratory air as a result of human activity and accidental contamination of mobile phase lines, filter sinkers, and bottles by leaving them exposed to the air is also considered to be a high risk activity and can lead to contaminant peak interferences.

	To mimic normal mobile-phase handling procedures, separate lots of mobile phase were prepared whilst wearing different types of glove. These included weighing and dilution of the salt, rinsing and filling of glass mobile phase bottles, and transferring the (pre-washed) solvent lines between mobile phase bottles. Results (Figure 2[a]) showed minimal additional contamination arising from use of (pre-washed) nitrile gloves compared to wearing the softer disposable nitrile gloves (Figure 2[b]), where significant extra contaminant peaks were observed, though in a much lower scale to when the whole glove was immersed. Leachable plasticizers from disposable gloves are likely to also be deposited on the hands and be transferred to anything touched after removal, including LC solvent lines and fittings, and as a result of this study the authors recommend that they should not be worn when handling anything in the LC flow path.

	Other sources of plasticizers such as plastic Pasteur pipettes, Parafilm, plastic tubing, and plastic weighing boats also gave rise to a significant increase in the number and level of contaminant peaks but at lower levels to those observed from gloves and dust. During this study it became evident that the size of some contaminant peaks present in the reference blank stock mobile phase solution increased significantly over time, presumably as a result of airborne contamination, microbial growth, or leaching from container, LC inlet filter sinker, solvent lines, or cap.

	Experiments were performed to determine and compare the baseline quality arising from a range of different grades of water and acetonitrile using the same method as described previously. It was observed that mobile phase prepared with commercial bottled HPLC gradient grade water produced lower quality baselines than mobile phase-prepared water from the laboratory water purification system. The apparent lower quality of commercial bottled water was consistent with previously published results (22). This could possibly be a result of the manufacturer’s purification system, insufficient container cleaning, cross-contamination with other solvents, microbes, environmental contamination, and leachates from storage vessels and caps. Laboratory water purification systems are capable of producing water with 5-50 ppb organic content, but only if they are regularly maintained and flushed. In line with previously published reports (24, 25) we also observed effects on baseline quality from different grades of acetonitrile. The baselines showed no significant difference between them in terms of the levels of observed contaminant peaks but the background UV response was higher with acetonitrile from some suppliers than from others. The LC-MS grade acetonitrile used in this study produced a generally lower baseline response compared to cheaper commercial HPLC grades. A possible explanation for this could be down the manufacturing process of acetonitrile because it is produced as a by-product from the large-scale production of acrylonitrile, which possesses a stronger chromphore than acetonitrile. We concluded that the highest available quality acetonitrile should be used when analyzing low-level solutes or when low UV detection wavelengths are required. 

	Different grades of ammonium acetate buffer salt were also compared. Results showed significant variations in baseline quality arising from the use of each batch. Surprisingly, the largest contaminant peaks were observed from the use of one of the highest-grade (99.999+ label purity) materials investigated. This buffer salt was supplied in a soft plastic screw-cap bottle, so it is possible that plasticizer leachates were responsible for these additional peaks. The level of contamination originating from the container could also possibly be exacerbated by scraping the sides of the container during weighing and by the deliquescent nature of ammonium acetate. 

	A range of extraction techniques were evaluated for their effectiveness in removing the contaminant peaks from identical lots of mobile phase prepared by deliberately adulterating clean aqueous ammonium acetate (containing 5% acetonitrile) with a range of different contamination sources. This was compared by summing the peak areas of those peaks eluting in the first half of the mobile phase gradient cycle (5-50% acetonitrile) and the second half of the cycle (50-95% acetonitrile) of each chromatogram separately so that any trends related to contaminant polarity could be observed. The degree of change in total contaminant peak response resulting from each treatment was compared (Table 1).

	Two commercial SPE cartridges packed with different stationary phases and a hydrophobic membrane filter disk were compared for their effectiveness at the removal of polar and non-polar contaminants from 500 mL of contaminated mobile phase. One cartridge was more effective in reducing the overall level of contaminant peaks but the other was more effective at removing non-polar contaminants (see Table 1). Despite thorough pre-rinsing of the cartridge, some additional signals (max peak height 5 mAU) were observed in the polar contaminant region of the chromatogram baseline after treatment with the latter cartridge possibly arising from leaching of UV absorbing components from stationary phase, frits, or column. Vacuum SPE, though simple to use, required significant additional processing by the operator and was found to be time-consuming and limited to small volumes. Using SPE devices on solvent inlet lines has been evaluated in our laboratories and found to be ineffective because channelling and cavitation affected the consistency of flow and effectiveness of contaminant removal. 

	The membrane filter disk SPE device was less effective at removing contaminants than the packed-bed SPE devices (also found in previously published results [40]), presumably as a result of reduced solute retention capacity from the reduced sorbent mass and bed depth. The flow-rate through the disk was very slow and the filtration process was time-consuming. This would be a limitation when large volumes need to be prepared. It is possible that repeating measurements using more diverse chemistries and more replicates may give a more accurate picture of the capabilities of off-line clean-up techniques for contaminant removal. However, the large volumes of mobile phase sometimes required and the use of off-line clean-up techniques leaves the mobile phase prone to subsequent contamination following the initial clean from airborne plasticizers, dust, and microbes made this unfavourable.

	SBSE was similarly tested for its effectiveness in the removal of contaminants. This method is highly attractive because of its simplicity and the fact that the mobile phase can be continually cleaned by spinning the stir-bar throughout the run time of the LC. Unfortunately for this application the approach was limited by the small stir-bar devices available. Larger capacity SBSE devices would be worthy of development for this particular application, perhaps using porous monoliths, but care would need to be taken to avoid phase-bleed. 

	On-line continuous clean-up techniques are advantageous because mobile phase purification occurs at or close to the point of use. Trap micro-columns made specifically for removal of contaminant peaks were placed between an aqueous pump and mixer on both HPLC and ultrahigh-pressure LC (UHPLC) systems and some contaminant removal was observed during the first few runs. However, contaminant breakthrough from the trap micro-columns occurred very quickly. As a result of their small size such columns have limited retention capacity and may be unsuitable for long sequence runs without continual flushing and regeneration as previously demonstrated (20, 22-24).

	In an attempt to increase the contaminant trapping capacity, the micro-column was replaced with larger scale (4.6 × 100 mm) turbulent flow chromatography (TFC columns). TFC adopts the use of large (typically 50-µm) porous particles allowing for high flow rates with low back pressure in the system (43), though with the flow-rates used in this study (1-3 mL/min) turbulent flow conditions would not be achieved. Two stationary phases were compared and both were found to be more efficient at removing the contaminant peaks than the lower-capacity micro-column trap (Table 1). The bonded silica C18 column was more effective at removing the contaminant peaks than the polymeric column where some small peaks were still observed in the polar region of the chromatogram. Clean up of the traps necessitated removal from the system, which is undesirable because handling mobile phase lines risks further system contamination.

	A limitation of siting a trap column between the mixer and the pump is that the trap column will contribute to the overall pump back pressure. Furthermore, this configuration is not possible with low-pressure mixing systems using proportioning valves because the aqueous mobile phase needs to be pumped through the phase to avoid cavitation and flow-channelling. Trying to aspirate the solvent through the trap would result in undesirable cavitation and flow-channelling. 

	To overcome the additional pressure limitation and system volume increase when using even larger trap columns, a new system was designed to allow the mobile phase to be recirculated through a semi-preparative scale trap column using an additional reagent delivery pump and low-pressure PEEK Tee-piece (Figure 3). Because the trap column was out of the flow-path of the LC column it was possible to use the larger trap columns without adversely affecting the chromatography. A semi-prep scale bonded silica C18 column (10 × 150 mm, 5-µm) was shown to remove 95% of the polar contaminant peaks and 100% of the non-polar contaminant peaks from the grossly contaminated mobile phase throughout 24 h of continuous operation (60 tests) leaving a clean baseline throughout the course of the run.

	To test the effectiveness of the recirculating mobile phase purification system in a commercially validated method it was applied to the purity testing of pharmaceutical samples containing oxycodone (OC). This method had proved troublesome in transfer between testing sites because of the presence of contaminant peaks originating from ammonium hydroxide and/or water used to prepare the mobile phase. The contaminant peaks interfered with the low level determination of OC and OC impurities by HPLC with UV detection. Contaminant peak profiles were found to vary from site to site and between different lots of reagent from the same supplier. This resulted in time wasted during investigations and the need to repeat analyses with fresh batches of mobile phase. Using a larger trap column the recirculating mobile phase clean-up system was successfully applied to this method. The interfering compounds from the mobile phase were removed and trouble-free operation was achieved throughout the two-week course of the study. Typical results from analysis of standards showing the baseline before and after cleaning are shown in Figure 4. Following completion of the study the trap was flushed with acetonitrile to remove the retained contaminants, thereby removing the need to handle the column or solvent lines.

	Successful method transfer between laboratories is at risk from contamination events arising from different reagent supplies and qualities and laboratory cleanliness and handling practices. The results obtained from these experiments are comparable to previous studies (17,20,â¨22-26), which showed that common laboratory consumables and the quality of reagents and solvents used in mobile phase preparation can have a significant impact on the formation of additional ghost peaks. Sub-optimal handling practices such as handling mobile phase lines whilst wearing plasticizer-laden gloves exacerbates the problem. 

	It is generally recommended that aqueous mobile phase should be freshly prepared to avoid microbial contamination of the water and contamination from airborne contaminants. However, it is very difficult to avoid contaminant peaks completely as a number of factors can cause them. The ubiquity of plasticizers and dust in modern laboratories makes it very difficult to completely avoid trace contamination of mobile phase occurring. It is recommended where possible to avoid the use of plastic materials in LC mobile phase and sample preparation particularly for high sensitivity LC-UV methods where solutes need to be measured at low concentrations. Typical laboratory air contains comparatively large quantities of dust and other airborne particles such as semi-volatile organic compounds (SVOCs) from air handling systems, which can readily adhere to surfaces and glassware and cause contamination. Therefore it is necessary to protect solutions and solvent lines from airborne contamination. Detergents used in glassware washing can also contribute to the contaminant peak profile unless residues are thoroughly rinsed from the glassware before use.

	The recirculating purification system that used large-scale HPLC columns proved to be the most effective of all the extraction techniques tested, removing almost all of the contaminant peaks for sustained periods of operation without compromise to system pressure or adding additional system volume. The columns were simply regenerated by flushing with acetonitrile after each run sequence or when required. The plumbing configuration using a separate reagent delivery pump is compatible with LC systems using both low and high pressure mixers without any compromise to system dead or dwell volume or pressure limitation as would be observed if the trap column were placed in-line. The mobile phase is constantly cleaned so any contaminants adsorbed from the atmosphere and other sources are continually removed before entering the analytical LC pump. In addition, large volumes of mobile phase can be cleaned through the trap column since the size of the column and stationary phase can be easily increased.

	The system does require an additional pump and plumbing, which, although easy to connect to any type of LC gradient pump, may prove prohibitive to routine use in some laboratories on the basis of cost. However, when faced with the need to measure multiple trace sample components without reference standards (as is common for pharmaceutical stability studies) and to transfer methods between sites operating with different quality water, solvent, and reagent supplies, it is more practical and cost-effective than resorting to MS for quantitative analysis. Furthermore, reducing the number of failed runs and investigations and speeding up data processing through routine generation of interference-free baselines easily offsets the extra cost. Application of this approach to cleaning of the organic solvent may enable the use of lower-cost solvent grades, though the effective purification of the aqueous mobile phase has the greatest positive impact on the quality of baselines generated in gradient reversed-phase LC.

	Even with ultra-clean handling processes it is very difficult to avoid trace contamination of the mobile phase in modern industrial laboratories. Contaminant peak interference in gradient LC is commonplace and even handling solvent lines whilst wearing gloves contributes to the contaminant peak profile. The identification and resolution of contaminant peaks from target analyte peaks of low-level impurities is time consuming and risks the quality and accuracy of purity measurements made in pharmaceutical release testing and stability studies. In this article a method is presented to eliminate the problem, allowing the operation of gradient LC-UV for extended periods of time. The system was simple to set up and scalable for use with different sizes of column, different column chemistries, and with any type of pump. Column regeneration and conditioning was also simple. Future gradient pump design could potentially incorporate reagent clean-up systems like this to help overcome the baseline problems caused by trace-level contaminants entering the pump and interfering with detection in LC-UV.

	The authors would like to thank Dr Frank David (RiChrom) for his donation of the SBSE stir-bars and Dr Hubert Quinn (Cohesive Technologies) for donation of the preparative scale TFC columns. Thanks to Dr Melissa Hanna-Brown and Dr Jonathan Beaman (Pfizer) and Dr Mark Parkin (King’s College, London) for their advice and assistance with manuscript review.

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Articles

Cover Story

This article presents a method for comparing the levels of baseline interference arising from common laboratory mobile phase contamination sources and assesses different approaches for removing dissolved contaminants to generate interference-free chromatogram baselines. The authors demonstrate that recirculating mobile phase through a semi-preparative scale column using a reagent delivery pump has advantages over previously published mobile phase decontamination methods.

Removal of Contaminant Peaks in Reversed-Phase Gradient Liquid Chromatography for Improved Detection of Pharmaceutical Impurities

	In gas chromatography (GC) split injection systems, as much as 99% or more of the injected sample never enters the column: It is released downstream of the inlet where it can encounter and possibly perturb precision gas control devices. The split-vent trap acts to prevent or at least moderate such effects. This month’s instalment addresses the operation and maintenance of split-vent traps.

	A plethora of gas chromatography (GC) injection techniques confront chromatographers in their daily work, and the related sample injection systems can be rather complex. Split, splitless, direct, programmed-temperature, headspace, thermal desorption, and more all serve the purpose of transferring analyte species of interest into the separation column, and each does so in ways uniquely suited to the nature of the sample and its matrix. Beyond simply moving sample molecules from outside the column to inside, injection techniques share some additional requirements. First, they should transfer sample accurately and repeatably. Of course, these two terms don’t have significant meaning until they are cast in the light of analytical needs that define accuracy and repeatability. Another desirable trait is less-frequent maintenance and tolerance of a larger number of injections without a loss of performance. Although unavoidable in some extreme cases, no one wants to disassemble an inlet every few samples.

	The primary consideration for achieving the best possible injection performance is a close match between the sample, the injection technique, and the column. The analysis of traces of methylene chloride in polycarbonate beads, for example, is much better matched to headspace sampling than to direct or splitless injection. Headspace sampling leaves polymer residues behind in a disposable vial, while whole-sample vaporizing techniques such as direct, split, or splitless injection would rapidly contaminate the inlet and inlet liner, necessitating frequent maintenance. On the other hand, determination of polynuclear aromatic hydrocarbons (PAHs) is not well-suited to headspace sampling because of the high temperatures needed to volatilize the target compounds. Instead, the higher temperature ranges of vaporizing injection makes it a much better choice. But then vaporized sample components that migrate beyond the inlet may cause other problems.

	Split injection helps match convenient and easy-to-dispense liquid sample sizes to the sample size and carrier-gas flow-rate requirements of open-tubular (capillary) columns. Liquid sample sizes of 0.5 µL or larger are best for handling by syringe; smaller injected sample volumes tend to compromise accuracy and repeatability because of evaporative and droplet-size effects. However, if the sample is relatively concentrated and it is all injected into the column, the total amount of solute may cause peak distortion because of overloading of the stationary phase. Split injection can reduce a too-large solute amount down to a volume the column can handle. Sample dilution is an alternative, but it may not be desirable as an extra sample preparation step or a source of possible contaminants. Split injection also reduces the amount of solvent in the column and avoids very large solvent peaks that might interfere with the separation of early-eluted solutes.

	Figure 1 illustrates the basics of split injection with back-pressure regulated pneumatic control. A constant mass-flow controller provides carrier gas to the split inlet at flow rates from as little as 10 mL/min up to 500 mL/min or greater. This split flow enters the inlet at point A and then flows into the inlet liner. A small amount of carrier-gas flow is diverted upwards, across the septum, and out the septum purge vent B. The main carrier-gas flow passes through the inlet liner and past the tip of the column at the split point C. Some of the carrier gas flows into the column entrance, while the rest continues out of the inlet liner and exits the inlet at split exit D. The internal seal isolates the incoming and outgoing carrier-gas flows.

	The split ratio, S, is the ratio of the flow through the inlet liner to the flow rate of the column, Fc. Both flows must be expressed at the same temperature and pressure, normally at standard conditions of 0 °C or 20 °C and 101 kPa. The liner flow is equal to the total flow through the inlet, Fs , minus the split-vent flow, Fv: 

	The split ratio would be 100:1, for example, with a column flow rate of 1.5 mL/min, a total split flow of 152.0 mL/min, and a split-vent flow of 2.0 mL/min.

	The split ratio refers to the ratio of the flows in equation 1 under static conditions and not to the actual fractional amount of sample that enters the column. The injection process itself is far from static. Upon entering the inlet liner, solvent and sample undergo rapid heating and expansion, which causes a pressure and flow impulse to develop and propagate along the liner in both directions. Note that this is not the “pressure pulse” of pressure-pulsed split-splitless injection, a technique that purposely increases the inlet pressure setpoint before injection and then reduces it back to normal after injection is complete. This higher initial inlet pressure can help narrow the injection pulse and control some of the effects of vaporizing injection.

	Column flow momentarily increases as the pressure impulse encounters the column entrance, but because of the relatively high restriction of the column and low restriction of the inlet liner and other inlet internal passages, the split flow will not react proportionately and the split ratio will deviate away from its initial quiescent value. The amount of each solute that enters the column is a complex function not only of the solvent, inlet temperature, injection amount, and inlet flow rates, but also of the liner packing and configuration, solute volatility, and injection speed. For these reasons, the split ratio should be taken as an approximation, and actual solute transfer into the column should be determined by careful calibration, along with recalibration upon modification of any aspects of the injection.

	After leaving the inlet at the split exit D, carrier gas passes through a split-vent trap, then through a back-pressure regulator, and finally is vented to the surrounding air. The split-vent trap serves two main functions. First, it acts as a buffer between the inlet and the back-pressure regulator that reduces the size of the injection pressure impulse that reaches the regulator. Second, it traps and holds less-volatile sample components that could otherwise contaminate the regulator and affect its operation permanently. An additional benefit is the trapping of toxic substances so they are not released into the laboratory atmosphere. An optional external split-vent trap, installed after the back-pressure regulator, at the external split-vent exit connection, can help ensure that toxics do not leave the instrument. Such external traps are useful if solvent exposure is a concern as well.

	The injection pressure pulse reaches the back-pressure regulator quickly, while sample splitting is still occurring inside the inlet. The regulator will sense the pulse and open itself up a bit to relieve the increased pressure. As the pressure pulse subsides, the regulator will head back to its previous static position. These fluctuations add to the carrier flow perturbations caused by injection and can increase uncertainty in the injected solute amounts as well as change the retention times of early-eluted solutes.

	By including the extra volume of the split-vent trap in the flow path, the extent of the pressure impulse is reduced, and it is spread out over a longer period. Fluctuations in the regulator are mediated and the inlet pneumatic system is made more stable. Using larger diameter tubing between the split exit D and the split-vent trap also helps mediate the pressure impulse.

	The initial injection pressure impulse propagates rapidly from the split-vent exit, at the speed of sound through the carrier gas and tubing. Shortly afterwards, moving at the split-vent flow rate, sample components and solvent start to move out of the inlet. In the absence of the split-vent trap, these would reach the back-pressure regulator within about 1 s after the start of injection and would further perturb its operation. Solvent vapour has a higher viscosity than carrier gas, and its presence at the back-pressure regulator would cause the regulator to open even further in compensation.

	The split-vent trap entrains solvent and sample components that are vented from the inlet, keeping them from entering the regulator immediately. With a split ratio of 100:1, this amounts to 99% of the total sample amount. Solvent will eventually break through the trap and be released slowly into the back-pressure regulator; over time most solvents will be purged from the trap. Less-volatile solutes will tend to accumulate, so that the back-pressure regulator is protected from the solutes condensing inside.

	Many chromatographers tend to neglect their split-vent traps, which are hidden underneath instrument panels and are easy to forget while troubleshooting or maintaining inlet systems. If normal maintenance procedures of replacing the inlet liner, internal seals, and septum do not remedy problems such as peak tailing, poor accuracy, and poor repeatability, or inability to achieve high split ratios, then the split-vent trap or connecting tubing may be at fault.

	Before replacing the split-vent trap, carefully check the split-vent and septum purge flow rates, preferably with an accurate digital flowmeter, and make sure there are no leaks around the inlet seals and septum. Leak checking is best done with an electronic leak sniffer and hydrogen or helium carrier gas. Using a surfactant solution on a cooled-off inlet will also find major leaks, but may then create a major headache because the solution can be drawn into the inlet by capillary action back through a leak.

	A blocked or restricted septum purge is a sure sign of a repeatedly overloaded inlet and the accompanying back-flash of sample up into the septum area. This problem is always accompanied by corresponding downstream contamination in the direction of the split-vent trap. While most common with splitless injection, low split ratios can yield the same effects. It may be necessary to replace the inlet nut assembly in this case, and if so be sure to replace the split exit tubing and split-vent trap as well. In extreme cases it may be necessary to remove the inlet entirely and solvent clean it.

	If the septum purge flow is normal, meaning about 2-5 mL/min, then 

	the problem may be a routine build-up of contamination in the split-vent trap. The trap may last for up to a year if injections are primarily clean volatile samples with little to no high-boiling residues. Samples that contain high levels of nonvolatiles, on the other hand, could necessitate trap replacement as often as monthly.

	With a replacement trap or trap cartridge on hand, proceed to remove the existing split-vent trap by following the instrument manufacturer’s procedures. With the trap removed, inspect the incoming tubing, the connections, and the trap itself for any visible residue. If found, such residue indicates the connecting tubing back to the inlet also needs replacement. This may be best performed with a service visit by an authorized technician. Consider modifying sample preparation in such cases so that as little high-boiling material as possible is injected.

	Be sure to leak-check the new connections and verify the correct split-vent flow rate, then try again to achieve good chromatographic accuracy and repeatability.

	Other injection systems also have vent lines that conduct sample components to the outside. Many headspace samplers, for example, vent a large portion of the sample. Thermal desorption systems usually have at least one split point as well. I remember one site visit where several headspace samplers were in use. Every one of them had a blocked vent exit tube, as evidenced by a complete lack of flow when the vent valve was activated manually. The sample was not particularly dirty, but the material under analysis, which needed an elevated thermostating temperature for good headspace results, had been condensing in the final short, unheated, section of vent line tubing for an extended period.

	Split injection vent traps are a necessary adjunct to high-performance gas chromatography with open-tubular columns. These traps protect carrier-gas regulation components in the short term from performance-degrading perturbations because of injection, and in the long term from accumulation of sample residues that can also affect performance. Split-vent traps also help prevent nonvolatile toxics from entering the laboratory atmosphere.

	Often, however, split-vent traps are not serviced and maintained as regularly as they should be. Residues can build up rapidly when analyzing many samples with heavy components, and contamination can sometimes extend back into split-vent connecting tubing. Careful attention to all of the inlet components, including split-vent and septum-purge elements, can significantly improve results and reliability.

 is a Senior Scientist at Serveron Corporation in Beaverton, Oregon, USA, and a member of LCGC Asia Pacific’s editorial advisory board. Direct correspondence about this column should be addressed to “GC Connections”, LCGC Asia Pacific, Hinderton Point, Lloyd Drive, Ellesmere Port, Cheshire, CH65 9HQ, UK, or email the editor-in-chief, Alasdair Matheson, at amatheson@advanstar.com

GC Connections

In gas chromatography (GC) split injection systems, as much as 99% or more of the injected sample never enters the column: It is released downstream of the inlet where it can encounter and possibly perturb precision gas control devices. The split-vent trap acts to prevent or at least moderate such effects. This month’s instalment addresses the operation and maintenance of split-vent traps.

Split-Vent Traps: GC’s Dirty Secret

Are your solvent costs too high? Here are some ideas on how to decrease the amount of mobile phase you use.

	In today’s laboratories, expense reduction has become a standard part of doing business. In some cases, the amount of money we spend on solvents for the mobile phase in liquid chromatography (LC) operations can be a significant part of the cost of analysis. In other cases, it can be less of a concern. For this month’s “LC Troubleshooting” discussion, I’d like to look at some of the options that may be appropriate ways to reduce the cost of mobile-phase solvents in your laboratory. There are three major ways to approach the problem: we can recycle all or part of the mobile phase and use it again; we can recover solvents through distillation of the waste mobile phase; and we can reduce the amount of mobile phase that we send through the column. I’ll consider each of these techniques below. Sometimes a combination of techniques may be used for further gains in cost reduction. Be aware that not all of these techniques may be available for your application. For example, direct recycling will work only for isocratic separations and not gradients.

	In my community, we have at-curb recycling for part of our household waste. This reduces the amount of waste piling up in the landfill and recovers certain materials, such as aluminium, steel, paper, and some plastics, for reuse. We are also encouraged to reduce the amount of waste we generate by reusing some materials and using less of others. When I return beverage containers to the grocery store, my deposit on these is returned. In some cases, the whole recycling process reduces my expenses, whereas in others it is done because it is the right thing to do from an environmental stewardship standpoint.

	When we consider the mobile phase used in LC operations, it may be possible to recycle all or some of the components, but even if it is possible, the economics of the process may not make practical sense. If your method is isocratic, the mobile-phase composition is constant, and you can directly recycle your mobile phase in many cases. This does not work for gradient methods, because the waste bottle will contain an “average” mixture of the mobile phase that is not directly useful.

	The simplest way to recycle the mobile phase is to direct the waste line from the detector directly back into the reservoir. This seems like a rash decision and many workers are reluctant to do this, even though it can be done successfully for many methods. “But”, you ask, “aren’t you contaminating the mobile phase with sample components that are eluted from the column?” Indeed you are, but let’s consider what is happening. Under isocratic conditions, the mobile-phase composition is constant, and even if you have a retained component of the mobile phase, it will not generate a peak in the chromatogram. For example, most ion-pairing reagents are chosen because they stick strongly to the column and build up a charge on the column packing. Under the steady state conditions of isocratic flow, you don’t see a peak for the ion-pairing reagent in the chromatogram. However, if you were to inject a sample that contained that same ion-pairing reagent - for example, hexane sulphonate - into a mobile phase that did not contain that compound, you might see a peak for the reagent if the detector were set to detect it. In other words, if a compound is introduced to the column at a steady state, its detector response will be constant and no peaks will be observed. This is exactly what happens if you direct the waste stream back into the reservoir.

	There are a few things to be aware of to minimize potential problems with direct recycling of mobile phase. First, you would like the mobile-phase composition to be constant and homogeneous. If you are continually adding sample waste components, the composition will not be truly constant, but if the dilution factor is sufficient, it will be constant from a practical standpoint. For example, if you have 1 L of mobile phase and you add 1 mL/min of waste to it while pumping 1 mL/min out of the reservoir, the volume will stay the same, but the composition will change very slowly over time. In this example, the already-diluted sample that is eluted from the column would be diluted 1000-fold more as it entered the reservoir. The background signal will gradually increase as more and more analyte is added to the reservoir, but this will generally not be noticeable over a few days’ time. To keep the solution as homogeneous as possible, place the reservoir on a stir plate and add a stir bar to keep it well mixed. Just as you need to be careful not to use a single preparation of mobile phase for too long without replacement so as to avoid evaporation, microbial growth, or other problems, you can’t use recycled mobile phase forever, either. You’ll have to determine how long you can use the mobile phase, but I’d suggest a maximum of 1-2 weeks. Generally, I prefer to make fresh buffers at least once a week even without recycling, and recycling will certainly not extend your current expiration policies.

	Direct recycling generally will be most successful when your analytes generate a large signal-to-noise ratio. It is most likely that any problems related to recycling will appear first for trace levels of analytes, such as with an impurity method, so direct recycling may not be practical for these methods.

	Because direct recycling gradually contaminates the mobile phase, an alternate recycling technique can be used, which I’ll call fractional recycling. If you consider a normal day’s LC operation in the laboratory, you put the mobile phase on the system and allow the column to equilibrate. Then you run a system suitability test, followed by some calibration standards, and finally run your samples for analysis, perhaps with occasional standards or check samples added. Of the total time of operation, for only a small fraction of the time is there a sample component being eluted from the column. If the fractions of the mobile phase that contain analytes and unwanted peaks, as well as the material eluted at the column void (solvent front), could be directed to waste, the remaining mobile-phase stream should be pure. If you put a two-way valve after the detector, you could watch the chromatogram, and when peaks were being eluted, you could direct the waste stream to a waste container; when no peaks were being eluted, you could direct the waste stream back to the sample reservoir. If you do this manually, of course, the labour costs will quickly eclipse any savings realized by reducing solvent consumption. What you need instead is a device that looks at the detector output and determines if a peak is present or not - this is what your data system does. By using a similar device to detect peaks and control the switching valve, you could have an automated recycler to replace the person running a manually controlled valve. Such a device is available from Spectrum Chromatography, called the S-3 HPLC Solvent Recycler. (I’m sure similar devices are available from other sources, as well.) The manufacturer claims that if you are able to recycle 25 L of high performance liquid chromatography (HPLC)-grade acetonitrile, you will pay for the device. If you use it, I’d suggest taking the same precautions I mentioned above for direct recycling: use a stir bar to keep the reservoir contents homogeneous, and don’t try to extend the expiration date of your mobile phase beyond current limits.

	As an alternate technique, you could use an automated switching valve and control it with the timed events output from your LC system. If you diverted the timed section of the chromatogram at the solvent front and perhaps where large peaks eluted, you would get the major advantages of fractional recycling without having to buy a specialized instrument. However, the commercial recyclers don’t cost much more than an automated switching valve, so it may not be worth your time to design your own device.

	When I started doing LC analyses in a pesticide residue laboratory in graduate school, our department distilled all our solvents to increase their purity. Today most of us simply purchase high-purity, HPLC-grade solvents. However, you could use distillation to recover the solvents from the waste mobile phase, perhaps with a spinning-band distillation column to get added purity. This would allow you to recover the organic solvent from an aqueous-organic mobile-phase waste stream that also contained buffers or other additives. Because distillation separates the organic solvent for recovery, it will work just as well for isocratic and gradient LC waste streams.

	At least one company, B/R Instrument Corp., offers several different distillation products to allow you to recover mobile-phase solvents. These recovered solvents can then be used to prepare fresh mobile phases. If I were to use one of these devices, I would like to be confident that it produces sufficiently pure solvents for LC analysis, especially if you are performing trace analysis with gradient elution. I’m sure that the noted supplier (or others) would be happy to discuss this concern with you.

	Perhaps the easiest way to conserve solvent is to reduce how much you use. You can make most of the changes discussed below and remain within the guidelines that the pharmacopoeias recommend. This means that all you need to do is verify that the new conditions give the same results as the original, document these results, and proceed with analysis. Your company may require additional procedures or documentation when such changes are made.

	As a general rule, if you keep the column chemistry, column plate number, and mobile-phase composition constant, you should obtain the same separation. Special attention needs to be paid to gradient methods, which may require compensating changes in the conditions, and for cases when instrument extra-column volume plays a significant role in band width. Next let’s look at three possible approaches to reduce mobile-phase use.

 Perhaps the simplest way to reduce solvent consumption is to reduce the column diameter. If the flow rate is adjusted for the change in column cross-sectional area, you should get the same separation and retention times with both isocratic and gradient methods. Take, for example, a method running on the most popular column, a 150 mm × 4.6 mm column, operated at a flow rate of 2.0 mL/min. We can generally reduce the column diameter to other popular sizes of 3.0 or 2.1 mm i.d. without excessive extracolumn band broadening. For the 3.0-mm column, the reduction in cross-sectional area is (3.0/4.6)2 = 0.4. To keep the same linear velocity of the mobile phase through the column, we need to reduce the flow rate by this same factor, so (2.0 mL/min × 0.4) = 0.8 mL/min. (As usual for these discussions, I’ve rounded or truncated calculated values, so your results may vary slightly if you repeat my calculations. Also, to avoid clutter I’ve not included units in some of the examples where they should be obvious.) This should give the same retention times as the original conditions, but with approximately 60% reduction in solvent used. In a similar manner we can calculate the effect of changing to a 2.1-mm i.d. column: (2.1/4.6)2 = 0.2. Apply this to the flow rate, (2.0 mL/min × 0.2) = 0.4 mL/min, and you’ll reduce your solvent consumption by 80%. You could use even smaller diameter columns, but if you reduce the diameter below 2.1 mm, it is likely that you’ll have excessive band broadening because of extra-column effects unless you modify the LC instrument.

 A reduction in column packing particle diameter (dp) will increase the column plate number inversely to the change in particle diameter. This, then, will allow you to use a shorter column for the same plate number. Take the example above for a 150-mm-long column packed with 5-µm dp particles. If we change to 3-µm particles, this will increase the plate number by 5/3 = 1.67-fold. For a constant plate number, we can reduce the column length by the same factor: (150 mm/1.67) = 90 mm. Because 90-mm columns aren’t commonly available, we’d round this length to 100 mm. This tells us that the 100-mm column packed with 3-µm dp particles would have approximately the same plate number as the 150-mm, 5-µm column. But because the length has been reduced by one-third, isocratic retention times would be reduced by one-third also, as would the method run time, so we’d save one-third of the solvent costs. In this example, the smaller particles will increase the pressure by (5/3)2 = 2.8-fold and the shorter length will reduce the pressure by 100/150 = 0.67-fold, resulting in an overall pressure increase of (2.8 × 0.67) = 1.8-fold. Many methods operate at pressures where this increase would still be within the instrument’s pressure capabilities. You could lower the flow rate to reduce the pressure; this would increase the retention times, but would not change the amount of solvent used per run. For example, reducing the flow rate 1.8-fold would mean operating at (2.0 mL/min /1.8) = 1.1 mL/min. A flow rate of 1.1 mL/min with a 100 mm column would give retention times of (0.67 × 1.8) = 1.2-fold larger than the original method, or a 20% increase in run time. An alternative would be to run at 1.5 mL/min for the same retention times and run time, but a 30% increase in pressure, which perhaps would be more acceptable. As discussed below, flow-rate changes beyond those used to generate constant linear velocity are appropriate for isocratic methods, but not for gradient methods without additional method adjustments.

Column Length and Diameter Plus Particle Size:

 By combining a change in column diameter with a change in the length and particle size, we can further reduce solvent consumption. Let’s consider that the original isocratic method used a 150 mm × 4.6 mm, 5-µm dp column operated at 2 mL/min and a run time of 10 min. This would use (2 mL/min × 10 min) = 20 mL of solvent per run. Reducing the column diameter to 2.1 mm was compensated for by reducing the flow rate to 0.4 mL/min for a solvent volume of (0.4 mL/min × 10 min) = 4 mL of solvent per run. If we combine this diameter change with the length and particle size change to a 100-mm, 3-µm column, we saw that this would reduce run times by 0.67. This would also reduce the total solvent used per run to (4 mL × 0.67) = 2.7 mL/run. We would now be using (2.7/20) ≈ 15% of the original solvent. I’ll leave you to calculate the change in pressure for this new condition.

	The above changes in length, diameter, particle size, and flow rate will not change the separation for isocratic methods, but more care needs to be taken for gradient methods. For gradients, it is necessary to keep the number of column volumes of solvent constant for each gradient segment when the conditions are adjusted, using the following equation:

	constant = (tGF)/Vm            [1]

	where tG is the time for each gradient segment (in minutes), F is the flow rate (in millilitres per minute), and Vm is the column volume (in millilitres). A reduction in Vm is proportional to the change in column length times the cross-sectional area (or diameter squared). This means that for two conditions (with subscripts 1 = original and 2 = new), we need to have

	(tG1F1)/(L1dc12) = (tG2 F2)/(L2dc22)           [2]

	For any given set of changes, as proposed above, we can determine the necessary gradient time adjustment by solving equation 2 for the new gradient time, tG2:

	tG2 = tG1 (F1/F2) (L2/L1) (dc2/dc1)2            [3]

	Let’s apply equation 3 to the above example, where we changed from a 150 mm × 4.6 mm, 5-µm dp column operated at 2.0 mL/min to a 100 mm × 2.1 mm, 3-µm dp column operated at 0.4 mL/min. Let’s further assume that the original gradient time, tG1, was 10 min. What is the new gradient time, tG2, that is required to get the same separation under the new conditions? Using equation 3, we get our answer: tG2 = 10 min (2.0/0.4) (100/150) (2.1/4.6)2 = 6.9 min

	The original method used (2.0 mL/min × 10 min) = 20 mL of solvent; the new conditions use (0.4 mL/min × 6.9 min) = 2.8 mL. Thus,the revised method would use (2.8/20) ≈ 15% of the original solvent, the same savings as for the isocratic method. Note that unless we change the gradient time as in equation 3, the separation (peak spacing) will change. For a multistep gradient, the time for each gradient step will need to be adjusted using equation 3. Additional method tweaking may be required when peaks are eluted early in the gradient, because the system dwell volume can be important for peak spacing of early peaks.

	The examples in this section assumed conventional LC conditions, but the same changes will apply equally for ultrahigh-pressure liquid chromatography (UHPLC) or changes from LC to UHPLC. Also, be aware that a smaller-volume column may require a reduction injection volume or injected sample mass to avoid volume- or mass-overload of the column.

	Most of us are driven by the economic factors in the laboratory, so be sure to look at all of the costs when applying any of the solvent-saving techniques discussed above. For example, if you have to buy a new piece of hardware for $1000 or more, how many litres of solvent need to be saved to break even (and don’t forget the cost of labour)?

	Usually the most practical and inexpensive way to reduce solvent consumption is by changing the column length, diameter, or particle size, as well as the flow rate. If you work in a regulated environment, such as pharmaceutical or environmental analysis, be sure that any changes are within the limits set by your company’s standard operating procedures (SOPs) and the regulatory guidelines. Documentation of such changes is essential for all applications; validation requirements will vary depending on the SOPs, the application, and the magnitude of the change involved.

 has been writing “LC Troubleshooting” for LCGC for more than 30 years. One of the industry’s most respected professionals, John is currently the Vice President of and a principal instructor for LC Resources in Lafayette, California. He is also a member of LCGC Asia Pacific’s editorial advisory board. Direct correspondence about this column should go to: “LC Troubleshooting”, LCGC Asia Pacific, Hinderton Point, Lloyd Drive, Ellesmere Port, CH65 9HQ, UK, or e-mail the editor-in-chief, Alasdair Matheson, at amatheson@advanstar.com

LC Troubleshooting

How to Reduce Mobile-Phase Consumption

, Supelco/Sigma Aldrich, Pennsylvania, USA.

	Hydrophilic-interaction chromatography (HILIC) is a complex system involving partition, polar, and ion-exchange interactions. Method development can be greatly facilitated by understanding the interactions that the different stationary phases provide and applying that knowledge to the separation task at hand. Through an understanding of the main differences of the set of analytes to be separated, a column or set of columns can be judiciously chosen to screen for effective retention and selectivity.  

	Analogous to reversed-phase chromatography, polar stationary phases used in hydrophilic-interaction chromatography (or HILIC as coined by Alpert [1]) provide different interactions that can be exploited by chromatographers to retain and separate various components of a mixture. Mechanisms of interaction in HILIC include partitioning, polar interactions, and ionic interactions (2). Partitioning involves the phase transfer of polar analytes from an organic-rich mobile phase into an adsorbed layer of water on the stationary phase. Polar interactions may then occur between the active surface or ligands of the stationary phase and ionic interactions may occur between charged analytes and oppositely charged moieties on the stationary phase. Stationary-phase chemistries can be designed to heighten or attenuate the various mechanisms and thus impart alternative retention and selectivity. For example, bare silica has been shown to adsorb water in the presence of an organic-rich mobile phase and thus provides the opportunity for an analyte to partition. Bare silica also exhibits negatively charged silanols at moderate pH conditions and above that may interact strongly with positively charged analytes (ion-exchange) (3). A penta-fluorophenyl phase has been shown to exhibit very little partition, yet provides a high degree of ion-exchange potential (4). On the other end of the spectrum, a pentahydroxyl stationary phase predominantly retains analytes though partition mechanisms because of the high degree of water adsorbed by the surface and shows relatively little ion-exchange capacity. Through an understanding of the interactions stationary-phase chemistries provide, one can choose the most appropriate blend of interactions that best complements a given separation challenge.

	HILIC is characterized by the use of a polar stationary phase and a relatively nonpolar mobile phase containing a small amount of water (1). The more polar water preferentially absorbs on the polar stationary phase providing an environment conducive to partitioning mechanisms. If a polar molecule, then, finds itself residing in the relatively nonpolar mobile phase it will tend to partition into the more desirable polar aqueous realm. Conditions are thus adjusted by chromatographers to induce both partitioning into and out of the aqueous layer. The system is pictorially shown in Figure 1. 

	Unfortunately, the situation is not that simple. By definition, polar molecules are interactive. Polar molecules are often ionic and thus may interact via ion-exchange mechanisms in addition to partitioning. At moderate pH and higher, silica-based phases exhibit negatively charged ionized silanols that may interact strongly with positively charged analytes. Negatively charged analytes may also be repulsed. Polar molecules may exhibit strong dipole moments or hydrogen bonding moieties that interact via adsorptive polar interactions. In HILIC, these highly interactive molecules are drawn very close to another polar, highly interactive surface making ion-exchange and polar interactions highly probable.

	Polar and ionic interactions are mechanisms that exist in the more familiar reversed-phase chromatography; however, these interactions are generally regarded as secondary and thus less important to partitioning. In HILIC, however, polar and ionic interactions are significant contributors to overall retention and selectivity and, in fact, are often the main mechanisms of interaction.

	Figure 2(a) shows a retention profile for the polar basic compound synephrine using two different HILIC stationary phases. In this figure, the retention is shown as a function of increased acetonitrile in the mobile phase and consequently the dilution of the buffer concentration. On both phases, the retention of synephrine is shown to increase at high acetonitrile content and thus both systems appear to be “HILIC”. The plot in Figure 2(b) shows the same experiment, but the buffer concentration of the mobile phase was kept constant as the organic modifier was increased. This approach has a dramatic effect on one column and virtually no effect on the other. What this demonstrates is that all retention under HILIC conditions is not necessarily because of partitioning mechanisms. On the bare silica phase, water is indeed adsorbing on the surface and thus providing the opportunity for the polar synephrine to partition. The retention is not impacted to a large extent by the change in buffer concentrations. On the pentafluorophenyl phase, however, there appears to be little or no water on the surface. Retention is therefore based on polar or ionic interactions and is highly influenced by the buffer concentration.

	The variables used to control and adjust partition and ionic interactions are very different. It is therefore paramount to have some understanding of the interactions that take place in a given system. It is also important to have a firm grasp on the interactions different stationary-phase chemistries provide to make sound choices during method development. As in reversed-phase chromatography, understanding retention mechanisms in HILIC can greatly facilitate method development and result in robust and rugged methods (5).

	For any analysis where the potential for ion-exchange exists, a good general practice is to evaluate the ion-exchange contribution to the overall retention and selectivity. For an ion-exchange process involving a singly charged analyte, the dependence of retention on the mobile-phase competing ion may be expressed as follows (6):

	log k = -log [C] + log βIEX          [1]

	where k is the capacity factor of the analyte, C is the concentration of the competing mobile-phase modifier, and βIEX is a constant for a given system that includes the phase ratio, ion-exchange capacity of the stationary phase, and the ion-exchange equilibrium constant. A plot of log k versus log[C] will thus yield a slope of -1 when ion exchange is solely responsible for the retention of the analyte, whereas the plot would yield a slope of 0 where ion exchange is not present.

	Figure 3 shows the results of such a study on Ascentis Express F5 (pentafluorophenyl), Ascentis Express OH5 (pentahydroxyl), and Ascentis Express HILIC (bare silica) stationary phases (all from Supelco) using ephedrine, a polar basic analyte, as the solute. On the pentafluorophenyl phase the resulting slope approaches a value of -1 indicating a dominant contribution to retention from ion-exchange. The pentahydroxyl phase, however, exhibits a slope nearing 0 indicating relatively little ion-exchange character. The bare silica phase shows a slope in the mid-range indicating a significant contribution from ion exchange, but also a significant contribution from other mechanisms. Because polar interactions are difficult to isolate, the “rest” of the retention can be deemed to be caused by a combination of polar and partitioning mechanisms. Table 1 provides a list of some common HILIC phases and the resulting ion-exchange contributions from a similar study using normetanephrine as a probe. 

	In addition, the table shows the retention of cytosine, a neutral analyte under the run conditions that is indicative of the relative partition and polar interaction contributions for each phase.

	The pentafluorophenyl and cyano phases both show relatively high ion-exchange contributions to the normetanephrine retention along with little partitioning or polar interactions as noted by the lack of cytosine retention. This finding is consistent with recent studies of Soukup and Jandera (7), where they used coulometric Karl-Fischer measurements to determine preferential uptake of water for different HILIC stationary phases. Simply stated, the results show that phases like cyano and pentafluorophenyl exhibit ion exchange, but relatively little partitioning. 

	On the other end of the spectrum lie the pentahydroxyl and BEH Amide (Waters) phases. Both stationary phases exhibit low ion-exchange and relatively high partition or polar interactions as demonstrated by the near zero slope values and the strong retention for cytosine, respectively. These findings are consistent with a study published by McCalley (3) under relatively high buffer concentrations.

	The bare silica phase showed moderate ion-exchange contributions along with moderate retention for cytosine. This is again consistent with the other works cited. The hybridized silica demonstrated low ion-exchange contributions, presumably because of the lower number of silanol groups present. The hybridized silica column did exhibit good retention for cytosine and thus represents a different relative contribution of ion-exchange and partition or polar interactions.

	Surprisingly, the Diamond Hydride phase (Cogent) showed moderately strong ion-exchange character. The Diamond Hydride phase is built on type-C silica that is largely populated with silicon-hydride (Si-H) groups instead of silanol groups (Si-OH) as found on traditional silica surfaces (8). With the majority of the surface rendered nonionic, low ion-exchange behaviour would be expected. The relatively strong ionic character observed remains unexplained, but is consistent with the findings of Bawazeer and colleagues (9). The Diamond Hydride phase exhibits moderate retention for cytosine and thus represents another useful blend of the various retention mechanisms.

	Also surprising were the relatively high ion-exchange contributions from both the Tosoh Amide (Tosoh Bioscience) and the Nucleoshell HILIC (zwitterionic) (Macherey Nagel) phases. Amide phases are generally characterized by low ion-exchange and high partition or polar interactions; however, based on the present study the Tosoh Amide phase appears to exhibit relatively high ion-exchange and high partitioning or polar contributions. Similarly, zwitterionic phases tend to have low ionic interactions and high partition or polar interactions, while the Nucleoshell column shows high ionic character and only moderate partitioning or polar mechanisms. These findings may be because of the conditions under which they are obtained, the manner in which the phases are bonded, or the chemical properties of the surfaces they are built on. Needless to say, one cannot conclude that all “amide”, “zwitterionic”, or especially “HILIC” phases provide the same blend of interactions.

Considerations for HILIC Method Development

	There are many different sets of analytes one may wish to apply to HILIC chromatography. A good starting point for method development is to identify the stationary phase with the appropriate mechanisms that best retain and differentiate the analytes. At a minimum, understanding the attributes of the stationary phase can be used to eliminate those phases that are inappropriate. For example, if a set of polar neutral molecules are to be separated, polar and partitioning mechanisms will be required whereas ionic interactions would be ineffective. Using a pentafluorophenyl stationary phase that primarily exhibits ion exchange would then be a wasted effort. The key to using any chromatographic mode is to have some idea of where the greatest differences lie with respect to the different analyte physiochemical attributes and invoke the most selective retention mechanisms by choosing the most appropriate phase or phases to screen.

	For method development in HILIC, a sound approach is to

		Understand the analyte properties. (Will they partition? Are they polar enough to partition into the aqueous phase? Will they ion-exchange? Are they charged, positively or negatively? How do they differ most, solubility (log P/log D) or degree of ionization (pKa values)?

		Choose the most appropriate stationary phase or phases that complement the largest differences in the analytes or, conversely, eliminate those that do not.

		Screen the chosen phases for the best selectivity.

		Optimize the separation using the most appropriate column chemistry.

	The development of a HILIC separation for a set of nucleosides is used to demonstrate the utility of this approach (10). Nucleosides in general are polar molecules and are weakly basic, thus falling under the category of “polar neutrals.” Cytidine, for example, exhibits a basic pKa value of 4.3 whereas its most acidic pKa is 13.5. Below a pH of 4.3 the compound would predominantly carry a positive charge, but under most HILIC conditions, the effective pH is such that the compound will be neutral. Figure 4 and Table 2 present the structures of the study analytes and pertinent physicochemical data, respectively. The set of compounds were screened using bare silica, pentahydroxyl, and pentafluorophenyl phases under several sets of HILIC conditions. As one would predict, only those phases exhibiting partition mechanisms proved useful.

	The majority of the nucleosides are neutral within the useful chromatographic pH window and thus cannot interact via ion exchange. To provide retention for the polar neutral molecules, the stationary phase must provide a partitioning mechanism. Both the pentahydroxyl and bare silica phases are polar enough to adsorb water onto their surfaces, thus enabling the potential for partitioning to take place. Figures 5 and 6 show the screening results for the pentahydroxyl and bare silica columns, respectively. Both phases provide good retention and selectivity for all of the probes. It is interesting to note that the last three compounds eluted show increased relative retention on the bare silica phase as compared to the pentahydroxyl phase. The retention of these late-eluted compounds also varies with pH on the bare silica phase, but their retention is relatively stable using the pentahydroxyl system. Both observations indicate significant ion exchange is occurring on the bare silica phase and demonstrates the limited ion exchange exhibited by the pentahydroxyl phase. The late eluted compounds were identified as 1-methyladenosine (basic pKa 6.1), 3-methycytidine (basic pKa 8.9), and the permanently positively charged 7-methylguanosine. Figure 7 shows the same experiments run on the pentafluorophenyl phase for demonstration purposes. As expected, little or no retention is observed for the polar neutral molecules because of the lack of partitioning provided by this particular phase chemistry. Again, it appears that a few of the nucleosides are charged enough to retain on the pentafluorophenyl phase via ion-exchange mechanisms. 

	The screening data indicated that the pentahydroxyl column using a pH of 5 provided the most selective and efficient starting point for further method development. These conditions were slightly refined to provide the useful separation shown in Figure 8. As per Table 1, column chemistries such as the Waters BEH HILIC and BEH Amide that show low ion-exchange interactions and significant partitioning might also be expected to produce similar retention and selectivity. This has indeed been found for the BEH Amide phase by Li and colleagues (11).

	Many examples of this approach can be found in the literature. For example ephedrine and related compounds were separated under liquid chromatography-mass spectrometry (LC-MS)-compatible conditions via primarily ion-exchange mechanisms on a pentafluorophenyl phase (12). For the separation of cathinone-derived designer drugs, both partition and polar interactions along with ionic mechanisms were required to resolve isobaric analytes (13).

	HILIC chromatography is a complex system involving partition, polar, and ion-exchange interactions. Method development can be greatly facilitated by understanding the interactions that the different stationary phases provide and applying that knowledge to the separation task at hand. A straightforward approach towards determining the relative ionâexchange contribution for various HILIC stationary phases has been described and can be used to classify HILIC columns by their predominant mechanisms of interaction. 

	The investigation demonstrated that diol/pentahydroxy and BEH Amide phases provide relatively little ionâexchange interactions and relatively high partitioning and polar interactions. Conversely, pentafluorophenyl and cyano phases exhibit primarily ion-exchange and relatively little partitioning character. Bare silica and several variations of HILIC columns are shown to provide different blends of ionâexchange, partitioning, and polar interactions. For a neutral set of molecules, such as the nucleosides, only partition can be expected to provide interactions that result in retention and selectivity. Indeed, the pentahydroxyl and the bare silica phase were shown to be useful and worth the time to investigate. Through an understanding of the main differences of the set of analytes to be separated, a column or set of columns can be judiciously chosen to screen for effective retention and selectivity. 

		D.V. McCalley in Hydrophilic Interaction Chromatography: A Guide for Practitioners, B.A. Olsen and B.W. Pack, Eds., (John Wiley & Sons, Hoboken, New Jersey, USA, 2013), pp. 1-41.	

		C.R. Aurand, H. Cramer, J. McKenzie, and D.S. Bell, 

		J.J. Pesek, M.T. Matyska, R.I. Boysen, Y. Yang, and M.T.W. Hearn, 

		S. Bawazeer, O.B. Sutcliffe, M.R. Euerby, S. Bawazeer, and D.G. Watson, 

		D.S. Bell and H. Cramer, Supelco Reporter 32.1, 6-8 (2014).	

		F. Li, J.A. Duan, D. Qian, S. Guo, Y. Ding, X. Liu, Y. Qian, Y. Peng, Y. Ren, and Y. Chen, 

		D.S. Bell, H.M. Cramer, and A.D. Jones, 

		X. Li, C.E. Uboh, L.R. Soma, Y. Liu, F. Guan, C.R. Aurand, D.S. Bell, Y. You, J. Chen, and G.A. Maylin, 

 is a manager in pharmaceutical and bioanalytical research at Sigma-Aldrich/Supelco. With a B.S. degree from SUNY Plattsburgh and a PhD in Analytical Chemistry from The Pennsylvania State University, Dave spent the first decade of his career within the pharmaceutical industry performing analytical method development using various forms of chromatography and electrophoresis. During the past 15 years, working directly in the chromatography industry, Dave has focused his efforts on the design, development, and application of stationary phases for use in HPLC and hyphenated techniques. In his current role at Supelco, Dave’s main focus has been to research, publish, and present on the topic of molecular interactions that contribute to retention and selectivity in an array of chromatographic processes. Direct correspondence to: dave.bell@sial.com

 is the editor of “Column Watch,” an analytical consultant, and a member of LCGC Asia Pacific’s editorial advisory board. Direct correspondence about this column should be addressed to “Column Watch”, LCGC Asia Pacific, Hinderton Point, Lloyd Drive, Ellesmere Port, Cheshire, CH65 9HQ, UK, or email the editor-inâchief, Alasdair Matheson, at amatheson@advanstar.com

Column Watch

Hydrophilic-interaction chromatography (HILIC) is a complex system involving partition, polar, and ion‑exchange interactions. Method development can be greatly facilitated by understanding the interactions that the different stationary phases provide and applying that knowledge to the separation task at hand. Through an understanding of the main differences of the set of analytes to be separated, a column or set of columns can be judiciously chosen to screen for effective retention and selectivity.

Retention and Selectivity of Stationary Phases Used in HILIC

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LCGC Asia Pacific-11-01-2015

Removal of Contaminant Peaks in Reversed-Phase Gradient Liquid Chromatography for Improved Detection of Pharmaceutical Impurities

Split-Vent Traps: GC’s Dirty Secret

How to Reduce Mobile-Phase Consumption

Retention and Selectivity of Stationary Phases Used in HILIC

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