Recent Developments in High-Performance Liquid Chromatography

Subscriber access provided by Milner Library | Illinois State University

Review

Recent Developments in HPLC Stationary Phases Thomas Chester Anal. Chem., Just Accepted Manuscript • Publication Date (Web): 02 Nov 2012 Downloaded from http://pubs.acs.org on November 5, 2012

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Analytical Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Analytical Chemistry

Recent Developments in HPLC Stationary Phases Thomas L. Chester* Department of Chemistry, University of Cincinnati, P.O. Box 210172, Cincinnati, OH 45221-0172, United States Experts and suppliers continue to make advances in the application of theory, capabilities, and practice of high performance liquid chromatography (HPLC). It is one of the most often-used analysis techniques and spans nearly every chemical application. Many users, even those with years of success operating an HPLC instrument and producing useful results, could benefit from recent knowledge of what is possible, what has changed, and what might be coming soon. There is little incentive to update or replace an old, reliable method when the cost and potential benefits of a replacement are uncertain. Awareness of what is available now and what is on the horizon is essential for the progressive practitioner.

We will focus primarily on the last two years of

developments, but will include several older reports of significance. The citations are not exhaustive, but are intended to represent recent work and advances. Although HPLC and supercritical fluid chromatography (SFC) are still practiced separately, the distinction is beginning to blur, and many stationary phases can be used in both techniques. So, we will include a few developments applicable to SFC. Also,

ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

achiral stationary phases will be our primary focus. Chiral stationary phase development was recently reviewed in this journal1. It seems like there are as many descriptions of types of innovations as there are authors who write about the innovation process. The simplest and most common innovations are continuous, that is, extrapolations or direct and predictable extensions from current practice. The biggest and most significant innovations, the real game changers, are discontinuous. They are also described as radical, revolutionary, or disruptive in their potential to improve or replace current practice. HPLC was built on numerous innovations. Instruments required the development of pumps, injectors, detectors, recorders, data processing, etc. Mobile phase innovations included blending solvents (to adjust strength), gradient elution, the use of buffers and additives, temperature control (which also affects the stationary phase chemistry, of course), and outlet pressure control (which enables the addition of condensed or compressed gases to the mobile phase). Stationary phase innovations included better adsorbents, bonded phases, pellicular particles, pore-size control, spherical particles, type B silica, chiral stationary phases, endcapping, polar endcapping, embedded polar groups, hybrid silica, polymeric bonding, etc. The subjective classification of each of these improvements as continuous or discontinuous is not as important as the actual capabilities they enable. SMALL PARTICLE ADVANCEMENTS AND LIMITATIONS Diameters of totally porous stationary phase particles decreased at a fairly regular rate throughout the history of HPLC2. Particles with diameters around 10 µm were widely used in the mid-1970s. The benefits of diameter reduction were entirely predictable, and


Page 2 of 41

Page 3 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


particle size (and plate height) diminished in concert with improvements in pumping, reductions in extra-column volume, and faster data acquisition until commercial particles reached about 2.5 µm in the 1990s. This was about the practical lower limit of particle size with 40-MPa (6000-psi) pumps, particularly if a cushion is desired in a method to allow a column to remain in use if the column resistance rises with age, or if one of the more viscous solvent systems (like water-methanol) is required. Monolothic columns emerged around 19963. The first commercial product appeared in 2000 and was disruptive: Compared to packed columns of the time, these monoliths had the potential to deliver significant plate height improvement and faster analyses, and they required substantially less pressure. For conventional packed columns to compete in efficiency and speed, the particle size had to be reduced below 2 µm, the delivery pressure had to be substantially increased, and the extra-column volumes shrunk. Thus was born ultra high performance liquid chromatography (UHPLC) and a new generation of instruments capable of utilizing sub-2-µm stationary phases. Today we have approached and perhaps reached the limit of increased performance via straightforward particle size reduction in UHPLC with totally porous stationary phase particles. This limit is due to frictional heating of the mobile phase, the resulting radial temperature gradients produced by the heating, and the consequential contributions to peak broadening from these radial gradients. Another limit is the difficulty of packing uniform beds with ever-smaller particles using current techniques. improvements may require one or more discontinuities. SLIP-FLOW CHROMATOGRAPHY


Additional


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 41

Perhaps the most intriguing new possibility is the use of slip-flow conditions in HPLC, as was recently demonstrated by Wei, Rogers, and Wirth4. Slip flow, fig. 1, is a wellknown phenomenon in MicroElectroMechanicalSystems (MEMS): the mobile phase velocity is zero at flow-channel walls in ordinary (no-slip or Hagen-Poiseuille) conditions, as in conventional HPLC, but reduction in the channel diameters eventually leads to slip-flow conditions where the mobile phase actually moves against the surface of the channel wall. The result is a large reduction in the pressure required to maintain the flow compared to the pressure extrapolated from no-slip conditions. Another benefit is a substantial reduction in peak broadening due to the narrowing of the range of flow velocities over the width of a flow channel; these velocity differences are a major source of broadening in conventional HPLC. The authors used 470-nm calcined silica colloidal particles that were slurry packed into 2.5-cm-long beds in 75-µm-diameter fused-silica tubes and then silylated. The authors showed chromatograms of florescent-labeled bovine serum albumin and of labeled monoclonal anti-prostate specific antigen, and they achieved over one million theoretical plates in about two minutes. Much work remains to turn this demonstration into practical capability. For example, there are no reports yet for performance with small solutes.

Convenient injection

providing extremely small initial bands is required. The authors used a fluorescence microscope and detected solute bands directly on the column, so post-column detection remains to be explored. In addition, practical column production techniques must be developed. However, if the potential is eventually realized in the workplace, this will be a truly disruptive innovation.


Page 5 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


We now turn our attention to recent advances in widely practiced technologies. CORE-SHELL PARTICLES Solid-core particles coated with a thin layer of stationary phase, originally called pellicular but today known as core-shell particles, got their start in the 1960s with the work of Horvath et al.5 and of Kirkland6. Commercial versions began appearing in their current format in 20077. Since then there have been about 150 reports, applications, and reviews in this rapidly growing area. These particles provide several benefits compared to totally porous packings8, 9. First, they have much better reduced plate heights; totally porous particles must be about onethird smaller in diameter than core-shell particles to provide comparable plate generation. There is some disagreement as to the reason behind the improved performance. It is widely claimed that the improvement is due to the limited solute penetration depth in the shell, thereby improving mass transfer, and that the narrow particle size distribution for most core-shell materials provides more uniform packed beds. However, Guiochon and Gritti report that the mass transfer improvement contributed by the limited shell thickness is negligible for small solutes (although large solutes eluted with fast mobile phase velocities benefit from the restricted diffusion depth), that improved efficiency arises primarily due to a reduction in eddy dispersion, and that the narrow particle size distribution is not responsible8. Tallarek et al. have extensively studied packing and mass transport10-15. The second benefit of core-shell columns is that they require one-half to one-third the pressure to operate than do columns of the same overall dimensions packed with totally porous particles providing the same plate number. Thus, with core-shell columns, high



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 41

plate numbers can be generated in short times, just as in UHPLC, but on systems with conventional pumping capability if adequate attention is paid to broadening contributions from injection, connections, and detection. And third, thermal conductivity is higher in core-shell columns due to the solid cores of the particles2,

16, 17

. The radial thermal gradients are lower than in columns of totally

porous particles of the same particle size pumped with the same mobile phase at the same velocity. Better heat dissipation allows the further reduction of core-shell particle size (and even better performance) before encountering mobile-phase heating problems. Commercial core-shell phases with particles as small as 1.3 µm are now offered. The reduction of particle size is driven by needs for smaller plate heights, faster analysis times, and higher plate number to achieve the separations required by users. Plate heights improve as the particle size is reduced, but at the cost of increasing the pressure necessary to provide the required flow rate.

Improved performance via

decreasing particle size can proceed as long as additional pressure is available (and all the components of the system can survive, and heating does not become limiting). However, if a higher plate count is required to solve a problem after the pressure limit is reached, then the particle size must be increased and the column lengthened to raise the plate number further18, 19. This comes at the cost of increased analysis time. Thus, we have seen two recent trends: 1) the development of smaller core-shell particles to improve plate height (and plates per second) and analysis time when higher pressure is available, and 2) the development of larger core-shell particles to provide higher plate number at the pressure limit. This is especially important to users of systems with “conventional” pressure capabilities who need more plates to solve a problem.


Page 7 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Core-shell columns with several different bonded-phase chemistries are available from a growing list of suppliers. Table 1 lists examples of the columns available at the time of this writing. In addition, work continues to develop new core-shell materials. Paek et al. introduced a thin layer of aluminum onto 2.7-µm core-shell particles, and then deposited a layer of carbon on the aluminum20. Efficiencies were about 5-times better than similar phases prepared on porous zirconia.

Liu et al., using a self-assembly technique, greatly

increased the carbon content of reversed phase materials compared to conventional procedures21. Li et al. grew nanometer sized zirconia crystals on the surface of silica microshperes22. Landon et al. prepared reversed-phase core-shell particles using carbon cores coated with poly(allylamine) and nanodiamond, and then functionalized the surface with C18 and C823. Their materials were stable at high pH and elevated temperature. Efficiency was good for lightly retained solutes, but reduced plate height was around 20 for well-retained compounds. Finally, pore sizes were initially in the 90-100-Å range for sub-3-µm core-shell particles, but applications for peptides, proteins, and other large solutes require phases with larger pores. Schuster et al. developed a core-shell stationary phase with 160-Å pores that is applicable to solutes up to 15,000 Da. Similar phases have now appeared commercially from other suppliers24. Characterization and performance have been investigated by several groups. Gritti et al. measured and related physical particle characteristics and performance25. Olah et al. compared mass transfer in core-shell columns and monoliths26. Fekete et al. compared performance for small and large solutes on 1.7-µm core-shell particles and found that high flow rates can be used to improve analysis times without a large loss in plate



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

number27. Gritti and Guiochon examined mass transfer kinetics in several core-shell materials28.

Ruta et al.29 and Dioszegi et al.30 also examined kinetic performance.

Various HPLC performance topics are discussed in several other papers28, 31-34. Lesellier examined the kinetic performance of core-shell stationary phases in SFC35. The scope of applications of core-shell stationary phases is essentially the same as for totally porous particles. For example, recent reports addressed lipids36, sulfanoamides37, and phenols and polyphenols38-43. Chirita et al. compared ion-pairing and reversed-phase approaches for the analysis of catecholamines using core-shell stationary phases. They found better results with the ion-pairing approach44. DeGrasse et al. separated shellfish toxins and reported that using a core-shell stationary phase compared to fully porous particles saved two-thirds of the analysis time45. Applications of core-shell columns in hydrophilic interaction chromatography (HILIC) are also appearing46-50. Because of the strong alignment of scope of core-shell and totally porous particles, we will not discuss applications of core-shell particles in otherwise conventional 1-d HPLC in further detail. There have been several reports of the use of core-shell stationary phases in 2-d HPLC51-53 where speed, particularly in the second dimension, is an important consideration. Chocholous et al. compared core-shell columns with monolithic columns for sequential-injection chromatography (SIC) and found that both can be used54. MONOLITHIC COLUMNS While core-shell phases are comparatively difficult to prepare, and progress is coming from relatively few experts and from specialized manufacturers, the situation is quite different for monoliths.

Their relative ease of preparation has empowered many


Page 8 of 41

Page 9 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


independent researchers to participate, and much progress has been made. There have been over 600 journal articles and 50 reviews since 2010. For readers new to the technology, a 2007 review by Guiochon provides history, perspective, and promise for the future of the technique55. Several other recent reviews are also recommended56-61. Commercial rods.

The first commercial monolithic column, called Chromolith

(Merck), appeared in 2000.

This was a 4.6-mm x 100-mm monolithic-silica rod,

functionalized with C18, and packaged in a poly(ether ether ketone) (PEEK) jacket with HPLC end fittings. This column attracted much attention due to its good efficiency (plate heights around 10 µm for favorable solutes—not bad at the time) and the very low pressure it required to generate mobile-phase flow. The low pressure was due to the high permeability of this monolith, so higher flow rates could be used to shorten analysis times compared to the typical 3.5-µm porous particles popular at the time. However, these monolithic columns suffered from radial non-uniformities in the rods that limited performance62. Also, many users considered the price to be outrageous compared to conventional columns. So, despite a great start, the first-generation silica-rod monoliths did not garner much additional attention after the introduction of UHPLC2. A recent change in the production process has improved the chromatographic performance, but this new generation of silica-rod columns requires about four-times more pressure than the earlier version63, 64. It will be interesting to see how well this generation is accepted by users. Inorganic Monoliths. Monoliths can be made from silica, from other inorganics (usually metal oxides), from organic starting materials, or as hybrids. The production of



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 41

silica monoliths was recently summarized by Trojer et al.65, Ghanem and Ikegami66, and El Deeb67. Monolith production from other oxides was reviewed by Gawel et al.68. Walsh et al. focused on non-silica inorganic monoliths69. Many of these have surface chemistries that are much different than silica and can provide different selectivities. Organic and Polymeric Monoliths.

The preparation, characterization, and

performance of organic or polymeric monoliths have been the subject of several reviews: Nischang et al. noted that porous polymer monoliths are useful in a variety of applications including liquid chromatography and microfluidic devices70. They discussed preparation, stability, and characterization. Separately, Nischang et al. examined the preparation of polymer monoliths with respect to mass transport and its influence on small solute separations71. They concluded that inhomogeneity of the monolith limits performance and that preparing monoliths with better small solute performance will be challenging. M. L. Lee’s group recently reviewed preparation and bed structure of capillary monoliths, compared these with conventional HPLC stationary phases, and examined performance for small solutes72, 73. Lee’s group also recently described preparing C18incorporated monoliths74, zwitterionic monoliths75, and cation-exchange monoliths76, 77, all based on polymethacrylates. Zhou et al. described the preparation of a poly(dimethylsiloxane)-modified monolith for use in reversed-phase separations78. Badaloni et al. made hexyl methacrylate and ethylene

glycol

dimethacrylate

monoliths

using

gamma

radiation

to

induce

polymerization in capillary tubes79. The authors then applied this to nano-HPLC/mass spectrometry where they found the columns to be stable.


Page 11 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Krajnc et al. reviewed recent applications of methacrylate monoliths with emphasis on understanding characterization and interaction mechanisms80. Hybrid Monoliths. Zhu and Row summarized the preparation and applications of hybrid silica-based and hybrid polymer-based monoliths and noted that these columns are finding application in HPLC81. Vujcic et al. reviewed organo-silica-hybrid monoliths and metal-oxide monoliths, particularly those of hafnium and zirconium82. They reported potential use in proteomics. Wu et al. discussed a one-pot synthesis and the use of polyhedral oligomeric silsesquioxane as a cross linker in preparing hybrid monoliths83. Wang et al. recently prepared a hybrid mixed-mode monolith with hydrophilic and strong anion-exchange interactions84. Jandera et al. made hybrid columns from conventional C18 or aminopropyl silica particles by first packing a capillary column with the particles and then polymerizing with methacrylate85. Perhaps not unexpectedly, the resulting columns had efficiency and permeability intermediate to those of conventional packed columns and monoliths. Selectivity. Monoliths can be made very selective by incorporating specific features into the surface during polymerization. Zhang et al. synthesized a hybrid monolith containing cyclodextrin86. Arrua et al. summarized recent work regarding synthesizing polymeric monoliths with incorporated nanomaterials87. Krenkova et al. also reviewed the use of nanostructure modifications to impart selectivity to monoliths88. Chambers et al. incorporated C60 fullerenes into a polymethacrylate monolith89.

They made a

butylmethacrylate monolith that provided more than 110,000 plates/m for benzene. When they used carbon nanotubes these authors achieved about one-third the efficiency they realized with the fullerenes90. Aqel et al. also prepared a nanotube-containing



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

monolith91.

Tong et al. incorporated carbon nanosheets into a polymethacrylate

monolith92. High selectivity can also be achieved by molecularly imprinting monolith surfaces. Tan et al. discussed preparation, characterization, and application of such materials in HPLC and sample preparation93.

Zheng et al. also reviewed molecular-imprinted

polymers with focus on developments and application in HPLC and in capillary electrochromatography94. He et al. prepared a silica monolith, then incorporated vinyl groups, and then injected a solution of methacrylic acid, ethylene dimethacrylate, sulfamethazine as a template molecule, and an ionic-liquid porogen95. The authors then evaluated the resulting column and found it selective for sulfamethazine in sulfonamide mixtures. Chen et al. imprinted a monolith to recognize dibenzoyl-D-tartaric acid96. Mu et al. used molecular crowding to enhance the effectiveness of imprinted monoliths97. Zhao et al. imprinted ketoprofen on a monolith using metal ions as a mediator98. Another approach involves immobilizing a protein on the surface of a stationary phase in order to impart desired selectivity.

Calleri et al. discussed the preparation and

applications of protein-doped sol-gel monoliths99. Such columns are useful for smallmolecule drug screening.

If cellular membrane proteins are immobilized, then the

resulting technique can be called cellular membrane chromatography or cellular membrane affinity chromatography100-102.

Calleri et al. reviewed several other

possibilities for changing the surface character of organic monoliths99. Tetala et al.103, Arrua et al.104, and Spross and Sinz105 also reviewed the use of monoliths in affinity chromatography.


Page 12 of 41

Page 13 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Pfaunmiller et al. prepared several polymeric monoliths incorporating human serum albumin106.

The authors determined the concentration of albumin in the various

monoliths they prepared, and then evaluated the best ones using R/S-warfarin and D/Ltryptophan as solutes. Wang et al. prepared SiO2/TiO2-composite monoliths, which they then used to enrich phosphopeptides via metal oxide affinity107.

Although no

chromatography was done, the potential as a new medium for phosphoproteomics is noteworthy. Zhang et al. developed monoliths containing immobilized Ti4+ or Zr4+ with affinity for phosphopeptides108.

Hou et al. also developed a silica monolith with

immobilized Ti4+ aimed at applications in phosphoproteomics109. Reichelt et al. prepared and used hydrophilic poly(ethyleneglycol)-based monoliths to purify ovalbumin110. Analytical applications are yet to come. There are also examples of monoliths containing copper111 and iron112. Monoliths in 2d-HPLC Systems. Morisaka et al. used an ion-exchange column in the first dimension and a reversed-phase monolith in the second dimension for the separation of proteins113.

Chen et al. used comprehensive 2d-HPLC/time-of-flight mass

spectrometry, with a monolith in the second dimension, to screen anti-tumor components from herbal medicines114. Applications and Other Uses. Readers who would prefer a focus on proteomics including the use of monoliths (rather than on monoliths with applications in proteomics) may find a review by Rozenbrand and van Bennekom covering eight years of developments through 2011 to be useful115. In addition, reports of recent work have appeared from Iwasaki et al.116, Morisaka et al.113, Vitorino et al.117, Wang et al.107, and



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Zhang et al.118.

Page 14 of 41

Kortz et al. provide a summary of using monoliths with mass

spectrometric detection with attention to metabolites and proteins119. Zheng et al. also recently reviewed the use of protein-modified monoliths with emphasis on chiral selection120. Wistuba121, Chankvetadze122, and Zhang et al.123 also summarized chiral separations using monoliths. Cis-diol compounds are important in many biological processes and are often present in low concentration.

Boronate-affinity chromatography provides useful selectivity for

isolating and determining these solutes. Li and Liu recently summarized work in this field including the use of monoliths124. Several reports described procedures to produce boronate-affinity monoliths with specificity for glycoproteins54, 125-136. Li et al. described a polymeric monolith containing a Wulff-type boronate128. Liu et al. prepared monoliths incorporating sulfonyl-substituted phenylboronic acid129.

Li et al. prepared a silica-

boronate hybrid monolith using 3-acrylamidophenylboronic acid as the boronate affinity agent130. The authors noted high hydrophilicity, high binding capacity, and resistance to base hydrolysis. Liu et al. prepared a restricted-access boronate affinity porous monolith as a mimic of protein A for the specific capture of antibodies131. Monoliths are finding use in sequential injection chromatography and multisyringe chromatography, which were reviewed by Fernandez et al.132. Koblova et al. looked at aspects of forming gradients in SIC133. There is growing interest in monoliths with hydrophilic surfaces for use in techniques such as HILIC.

Jiang et al.134 and Gunasena and El Rassi135 reviewed monolithic

columns for HILIC applications. Incorporating ionic groups is also possible, as was reviewed by Nordborg et al.136, and these can be used in HILIC in various ways or as ion-


Page 15 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


exchange or ion-chromatography media. Zhang et al. reported a one-pot preparation of a strong cation-exchange monolith118. Yang et al. synthesized strong cation exchange monoliths137,

138

. Lv et al. described making a zwitterionic monolith with which they

separated nucleosides and peptides with good efficiency139-140. Guerrouache et al. also reported a zwitterionic monolith141. There have been several reports of silica-based monoliths for HILIC84,

134, 142

and of

hydrophilic polymer-base monoliths143-145. Sherikova and Jandera examined the effects of operational parameters for capillary zwitterionic-methacrylate-based monoliths146. They found excellent thermal stablity and good separations using high ionic strengths. Lin et al. described a one-pot preparation of a zwitterionic hybrid monolith147. Hydrophilic Interaction Chromatography and Related Techniques HILIC is another area of high enthusiasm. Several recent reviews provide the history, fundamentals, and perspective that will be important to new users148-154. Understanding and predicting retention in HILIC has been challenging, and Gika et al. recently investigated several retention models for gradient-elution HILIC155.

Creek et al.

developed a retention-time-prediction model and applied it to metabolite identification156. Stationary phases.

Development of HILIC stationary phases is occurring both

commercially and independently.

The need for different selectivity and better

performance drives the interest in stationary phase development. Bare silica can be used in HILIC, but surface modification provides a wider range of behavior and selectivity. Silica-based stationary phases were reviewed by Qui et al.157. Ray et al. recently described a peptide-modified silica for HILIC158. Li et al. made a polar silica phase containing poly-L-lysine159. This phase exhibited both reversed-phase



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 41

and HILIC character depending on the mobile phase composition. Li et al. also reported making a polysaccharide-modified silica160. Armstrong et al. have developed and investigated silica phases modified with cyclofructans. First they covalently attached cyclofructan 6 to silica161. They tested with several classes of polar analytes, examined the effects of several operational parameters, and concluded that the phase has very broad applicability to HILIC separations. Armstrong et al. later reported another new phase using sulfonated cyclofructan 6162. They followed with a more extensive investigation comparing both of these phases with bare silica and comparing intermolecular interactions163.

They concluded that these

surface modifications provided substantial performance improvements compared to bare silica. Armstrong et al. also reported making a stationary phase by bonding a diphenylphosphoniumpropylsulfonate to silica164. This phase was zwitterionic. The authors reported that this phase provided better performance than either bare silica column or a commercial zwitterionic column for the separation of several classes of solutes.

Li et al. modified silica by immobilizing imidazoline to provide both

hydrophobic and hydrophilic characteristics165. This phase was zwitterionic at the basic end of its pH range (4-9). Guo et al. made a zwitterionic phase by bonding L-azido lysine to silica166. Kotoni et al. modified silica to make two urea-type stationary phases, one with –NH2 groups and one without167. Fractions of each of these phases were then subjected to silanization to make four phases in all. These were evaluated by separating polyols, sugars, aromatic acids, nucleobases, and nucleosides. Schuster and Lindner developed


Page 17 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


stationary phases by browning aminopropyl silica with a Maillard process168.

The

resulting “chocolate” phases, named for their color, exhibited mixed-mode retention. Dai et al. used atom-transfer polymerization to make a polystyrene-based phase169. Huang et al. developed a glycosyl amino acid phase with glycopeptide selectivity170. Li et al. used carbon nanoparticles made from nitric-acid-refluxed corn soot as stationary phase, and they separated several polar solutes with HILIC conditions171. Kalafut et al. deposited carbon on a zirconia surface to produce a stationary phase with HILIC behavior172. Kawachi et al. characterized 14 commercial HILIC phases on the basis of hydrophilicity, the hydrophilic-hydrophobic selectivity, regio and configurational selectivity, shape selectivity, electrostatic interactions, and acid-base properties173. HILIC phases in SFC and related techniques. Polar stationary phases are most often used in SFC, and many HILIC phases work well174-176. West et al. examined eleven commercial HILIC stationary phases with 146 test solutes using SFC conditions176. The authors compared and classified retention using quantitative structure-retention relationships and principal component analysis, and evaluated peak shapes. Stationary phases fell into three groups according to their hydrogen bonding behavior. In this work, the mobile phase excluded water, so the retention mechanism was not HILIC. Starting with acetonitrile/aqueous-buffer or alcohol/aqueous-buffer mixtures and adding CO2, HILIC-like separations have been reported using enhanced-fluidity conditions by several groups. dos Santos Pereira et al. added CO2 in varying ratios to acetonitrile/aqueous buffer and ethanol/aqueous buffer mixtures177.

Only one

temperature and one pressure were used, and the organic-solvent/water ratio was fixed.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Other work was reported by Olesik et al.178-179. Philibert and Olesik examined retention for a fixed ratio of methanol and water while varying the amount of CO2 in the mobile phase179. From an SFC starting point, Taylor recently reviewed the use of water as an additive180. More specifically, Ashraf-Khorassani and Taylor used water added to various alcoholmodified CO2 mobile phases for the separation of several nucleobases on three widely utilized polar stationary phases181. Patel et al. used water as an additive in SFC along with ion-pairing agents182. Are any of these separations actually HILIC?

It is well established that surface

excesses of some mobile phase components commonly exist in SFC processes. The incorporation of water into a mobile phase combined with the use of a polar stationary phase is certainly necessary for HILIC, but is it sufficient? Alpert’s description of HILIC requires not just some water in the mobile phase but also a surface excess of water183. Determining if or when a surface excess of water exists, investigating how it changes with conditions, and examining retention behavior as a function of all adjustable parameters (including water concentration in the mobile phase, water/organic ratio, CO2 concentration, temperature, pressure, etc.) is necessary to discern the retention mechanism, to direct further research, and to develop new capabilities and useful applications. HILIC behavior, even with CO2 present in the mobile phase, is part of a continuum of chromatographic behavior including both conventional reversed-phase chromatography and conventional SFC; the best conditions for solving a particular separation problem is situated somewhere in this continuum184.


Page 18 of 41

Page 19 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


HILIC Applications.

New HILIC phases provide different selectivity and new

possibilities for separations, so we will take a brief look at applications of some of these new phases. Schiesel et al. used HILIC in investigating means to profile nutritional infusion solutions containing a variety of components185. HILIC has recently been used for determining carbohydrates including oligosaccharides186-191. Li et al. examined lowmw heparins using HILIC/ESI-Fourier-transform-MS192.

Walker et al. compared

reversed-phase and HILIC approaches to the analysis of derivatized N-linked glycans193. In seeking a single separation platform to support both proteomics and glycomics, they concluded that reversed-phase out-performs HILIC for their derivatives. There are several reports regarding analyses of nucleosides, nucleotides, etc.194-198. Lin et al. profiled urinary metabolites and found biomarkers for bladder cancer and for kidney cancer199. Stertz et al. used HILIC-MS/MS to show that 1-hydroxybutene-2-yl mercapturic acid is a good biomarker for humans as a screen for exposure to 1,3butadiene, a carcinogen200. Spagou et al. conducted metabolic profiling of rat urine in a hepatotoxicity study of galactosamine and the protective effect of glycine201. Several other reports of interest involve metabolites202-215. In proteomics, Di Palma et al. developed and applied a strategy requiring 100-timessmaller samples216-217. The proteome coverage was comparable to earlier methods. Zhao et al. described their development of a 2-d HILIC—reversed-phase/MS approach for rapid proteomic analyses218. Garbis et al. extended a multidimensional approach for proteome analysis of human serum219.

They combined size exclusion, HILIC, and

reversed-phase separations with detection by nanoelectrospray-ionization-tandem mass spectrometry. With this, they identified nearly 2000 proteins spanning 12 orders of



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

magnitude in concentration, and 375 phosphoproteins.

Page 20 of 41

Zarei et al. discussed the

application of HILIC and related techniques in phosphoproteomics220. Engholm-Keller et al. reported a strategy, involving HILIC, for phosphoproteomics in which they identified 4700 phosphopeptides221. Kaji et al. 222 and Wu et al. 223 also examine proteomics. MIXED-MODE AND ADJUSTABLE SELECTIVITY The boundary between HILIC and mixed-mode columns is very fuzzy. The bonded phases on HILIC columns are polar and hydrophilic. The addition of charged groups to the bonded phase increases hydrophilicity and adds the possibility of electrostatic interactions with charged solutes through both repulsive effects (as in electrostatic repulsion hydrophilic interaction chromatography, ERLIC) and ion exchange.

For

example, adding positive charges to the stationary phase can improve the peak shape of ionized bases at the expense of peak shape of acids224. Zwitterionic stationary phases affect the orientation of approaching zwitterionic solutes, such as peptides225.

So,

multiple interactions are possible. If the stationary phase is hydrophobic but contains ion-exchange groups, then reversed-phase and ion-exchange mechanisms can coexist. Yang and Geng recently reviewed mixed-mode chromatography in the context of biopolymer analysis226. The presence of mixed modes certainly adds complexity to method development, but it also adds incredible power, particularly adjustable selectivity, that can make method development quite easy. For example, consider the separation of acetaminophen (also called paracetamol) and its impurity, 4-aminophenol. (This is an important analysis problem in the production of pain-reliever products containing acetaminophen.)


Page 21 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


4-aminophenol is so polar that it cannot be adequately retained in a reversed-phase separation at neutral pH. This solute is cationic and even less retained with acidic conditions. Even if slight retention is achieved by reversed-phase for standards, polar excipients (such as flavorants, preservatives, etc.) in products also often elute early and interfere with the 4-aminophenol peak in actual samples. Switching from reversed-phase to HILIC provides different selectivity and higher retention of polar solutes, but even more control of both retention and selectivity is possible using a mixed-mode approach.

Calinescu et al., fig. 2, used a reversed-

phase/cation-exchange mixed-mode separation to retain the cationic form of 4aminophenol while not greatly affecting the reversed-phase retention of acetaminophen and other nonionic solutes present227. This kind of mixed-mode separation will allow a cationic solute to be positioned with great control into the valleys between the nonionic solutes, thus providing flexibility and adjustable selectivity to accommodate additional nonionic solutes that may also be present in commercial products. This control is accomplished by adjusting pH (if near the pKa of either the ionizable solute or the column) or mobile-phase concentrations of displacing cations with little effect on the nonionic solutes. Conversely, changing the organic modifier concentration affects the nonionic solutes in the usual reversed-phase manner with little effect on the ionic solutes. These numerous control possibilities also enable the expanded use of gradients in pH228, ionic strength, temperature, etc. in addition to ordinary modifier and flow gradients. Thus, the possibility of changing selectivity not only during method development but also during a chromatogram is possible. These capabilities enable a skilled worker,



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 41

empowered with knowledge of the chemistries of the solutes and of the stationary phase, to quickly and easily develop an alternative to a nearly impossible reversed-phase separation by using a mixed-mode approach. Mixed-Mode Stationary Phases.

Commercial mixed-mode phases have been

available for several years but have not been as widely adopted by users as is warranted by their potential.

Initial resistance was mainly due to lack of batch-to-batch

reproducibility for some of these phases and by serious errors and omissions in descriptions of how these phases behave. Progress has been made in reproducibility, particularly as more suppliers get involved, but technical information enabling informed decisions for method development still trails. Laemmerhofer et al. examined a silica-based, reversed-phase—weak-anion-exchange material incorporating alkyl groups with embedded thioethers and amides229.

The

functionalities of this phase, depending on both the solutes and conditions, could provide additional attractive or repulsive forces to retention. Zhang and Carr synthesized a silicabased, mixed-mode phase involving a polystyrene network with some carboxylate groups230. They also reported that gradients in organic modifier concentration, pH, and mobile-phase ionic constituents are possible, and mass-spec-compatible acids may be used.

Bicker et al. synthesized several urea-modified-silica stationary phases and

evaluated them under HILIC conditions231.

They observed a variety of retention

behaviors, depending on the properties of solutes.

Zhao et al. synthesized a

tetraazacalix[2]arene[2]triazine-modified silica that had mixed mode behavior232. Borges

et

al.

synthesized

and

characterized

a

thermally

immobilized

poly(dimethylsiloxane) phase bound to silica particles233. The stationary phase provided


Page 23 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


a reversed-phase mechanism with acidic mobile phases and mixed-mode reversedphase/cation-exchange with neutral mobile phase. Abood et al. made a hexylacrylatebased mixed-mode monolith234.

Zhang et al. grafted N-methylimidazolium to a

monolithic silica to create a stationary phase with multiple interactions235. Other mixed-mode combinations are possible, such as RP-HILIC236, exclusion/ion-exchange, etc.

237

, size-

Liang et al. combined C18 and C8 character with

chloropropyl character onto silica and applied it to alkaloid and glycoprotein separations238, 239. CONCLUSIONS There clearly has been much recent progress in HPLC stationary phases, and there is huge potential for further improvements.

Yet the number of academic researchers

developing chromatography technology is in decline, the number of new graduates with chromatography expertise (not just experience) is also declining, and the illusion of turnkey solutions from the suppliers reinforces the perception that HPLC is mature, easy to implement, and can be learned by attending a short course.

Sadly, technical

information on commercial stationary phases is often lacking. Rather than dumbingdown the chemistry for the least-capable users, suppliers should provide accurate information on the physical and chemical characteristics of stationary phases. Users should strive to increase expertise and fundamental understanding as new capabilities emerge.

Informed users could more effectively develop clever, cost-effective

separations, improve capabilities and efficiencies of their businesses, and even more effectively utilize innovations as they appear. Imagine the impact of doing HPLC better.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 1. Comparison of Hagen-Poiseuille flow (non-slip, as in conventional HPLC) and slip flow. Arrows in the channels are velocity vectors. Notice that the velocity is zero at the walls with Hagen-Poiseuille conditions but is moving at the walls with slip-flow conditions. Also note the reduction in the range of velocities with slip-flow conditions. Reprinted with permission from ref 4. Copyright 2012 American Chemical Society.


Page 24 of 41

Page 25 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 2. Separation of hydroquinone (1), acetaminophen (2), p-benzoquinone (3), 4aminophenol (4), 4-nitrophenol (5), and 4’-chloroacetanilide (6) on a Hypersil Duet C18/SCX column (250 x 4.6 mm, 5 µm) by gradient elution between phosphate buffer (pH = 4.88) and methanol. At this pH, 4-aminophenol is mostly cationic, and all the remaining solutes are uncharged. What is important to note here is the retention of 4aminophenol, its selectivity compared to acetaminophen, and the potential for this mixedmode approach to retain strongly hydrophilic ions and to adjustably position them among neutral solutes retained by a reversed-phase mechanism.

Reprinted from ref 227,

copyright 2012 The Author, by permission of Oxford University Press and I. A. Badea.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 41

Table 1. Examples of Commercial Core-Shell Stationary Phases Supplier

Product name

Advanced Material Technology

Agilent

MachereyNagel Phenomenex

Sigma–Aldrich

Sunniest Thermo Scientific

Halo

Particle diameter (µm) 2.7

Shell thickness (µm) 0.50

Halo Peptide-ES 160 (Å) Poroshell 300 (Å)

2.7 5

0.50 0.25

Poroshell 120 (Å)

2.7

0.50

Nucleoshell

2.7

0.5

Kinetex

5 2.6 1.7

N/A 0.35 0.23

1.3 3.6 1.7 3.6 2.7

N/A 0.2 0.22 0.5 0.50

Ascentis Express 2.7 Peptide-ES (160 Å) SunShell 2.6 Accucore 2.6

0.50

Aeris Widepore (200 Å) Aeris Peptide (100 Å) Aeris Peptide (100 Å) Ascentis Express

0.5 0.50


Surface chemistries

C18, C8, HILIC, RPamide, phenylhexyl, pentafluorophenyl, penta HILIC C18 SB-C18, SB-C8, SBC3, SB-CN EC-C18, EC-C8, Phenyl-hexyl, SB-C18, SB-C8, SB-Aq, BounsRP, HILIC RP-18, HILIC C18, XB-C18, HILIC, pentafluorophenyl C18, XB-C18, C8, HILIC, pentafluorophenyl C18 XB-C18, XB-C8, C4

C18, C8, HILIC, RPamide, phenylhexyl, pentafluorophenyl, pentahydroxy C18 C18 C18, aQ, RP-MS, HILIC, phenylhexyl, pentafluorophenyl

Page 27 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


AUTHOR INFORMATION Corresponding Author *E-mail: [email protected], Phone: +1-513-310-7375 BIOGRAPHY Thomas L. Chester received his Ph.D. degree from the University of Florida in 1976 under the direction of James D. Winefordner. He then moved to Procter & Gamble’s Miami Valley Laboratories where he served as Section Head and later Research Fellow in the Corporate Research Division. He retired from P&G in 2007 and is currently Adjunct Research Professor at the University of Cincinnati.

His research interests

include HPLC, modeling and optimization of separation methods, and the expansion of scope and the unification of separation techniques.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

References (1) Ward, T. J.; Ward, K. D. Anal. Chem. 2012, 84, 626-635. (2) Gritti, F.; Guiochon, G. J. Chromatogr., A 2012, 1228, 2-19. (3) Minakuchi, H.; Nakanishi, K.; Soga, N.; Ishizuka, N.; Tanaka, N. Anal. Chem. 1996, 68, 3498-3501. (4) Wei, B.; Rogers, B. J.; Wirth, M. J. J. Am. Chem. Soc. 2012, 134, 10780−10782. (5) Horvath, Cs.; Lipsky, S.;R.; J. Chromatogr. Sci. 1969, 7, 109 (6) Kirkland, J.;J. Anal. Chem. 1969, 41, 218. (7) Kirkland, J. J.; Langlois, T.; DeStefano, J. Am. Lab. (Shelton, CT, U. S.) 2007, 39, 18-21. (8) Guiochon, G.; Gritti, F. J. Chromatogr., A, 2011, 1218, 1915–1938. (9) Gritti, F.; Guiochon, G. LCGC North Am. 2012, 30, 586. (10) Khirevich, S.; Daneyko, A.; Hoeltzel, A.; Seidel-Morgenstern, A.; Tallarek, U. J. Chromatogr., A, 2010, 1217, 4713-4722. (11) Daneyko, A.; Holtzel, A.; Khirevich, S.; Tallarek, U. Anal. Chem. 2011, 83, 3903-3910. (12)

Bruns, S.; Tallarek, U. J. Chromatogr., A 2011, 1218, 1849-1860.

(13) Bruns, S.; Hara, T.; Smarsly, B. M.; Tallarek, U. J. Chromatogr., A 2011 1218, 5187-5194. (14) Khirevich, S.; Höltzel, A.; Daneyko, A.; Seidel-Morgenstern, A.; Tallarek, U. J. Chromatogr., A 2011, 1218, 6489-6497. (15) Daneyko, A.; Khirevich, S.; Höltzel, A.; Seidel-Morgenstern, A.; Tallarek, U. J. Chromatogr., A 2011, 1218, 8231–8248. (16) Kostka, J.; Gritti, F.; Kaczmarski, K.; Guiochon, G. J. Chromatogr., A 2010, 1217, 4704. (17) Kostka, J.; Gritti, F.; Kaczmarski, K.; Guiochon, G. J. Chromatogr., A 2011 1218, 5449. (18)

Knox, J. H.; Saleem, M. J. Chromatogr. Sci., 1969, 7, 614.

(19) DeStefano, J. J.; Schuster, S. A.; Lawhorn, J. M.; Kirkland, J. J. J. Chromatogr., A, 2012, 1258, 76– 83. (20) Paek, C.; Huang, Y.; Filgueira, M. R.; McCormick, A. V.; Carr, P. W. J. Chromatogr., A 2012, 1229, 129-139.


Page 28 of 41

Page 29 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(21) Liu, Q.; Wang, L.-T.; Dong, S.-Q.; Zhang, Zh.-X.; Zhao, L. J. Inorg. Organomet. Polym. Mater. 2011, 21, 941-945. (22)

Li, J.; Zhang, H.-F.; Shi, Y.-P. Anal. Sci. 2011, 27, 447-451.

(23) Wiest, L. A.; Jensen, D. S.; Hung, C.-H.; Olsen, R. E.; Davis, R. C.; Vail, M. A.; Dadson, A. E.; Nesterenko, P. N.; Linford. Matthew R. Anal. Chem. 2011, 83, 5488–5501. (24) Schuster, S. A.; Wagner, B. M.; Boyes, B. E.; Kirkland, J. J. J. Chromatogr. Sci. 2010, 48, 566-571. (25) Gritti, F.; Leonardisa, I.; Abiaa, J.; Guiochon, G. J. Chromatogr., A, 2010, 1217, 3819–3843. (26) Olah, E.; Fekete, S.; Fekete, J.; Ganzler, K. From J. Chromatogr., A 2010, 1217, 3642-3653. (27) 490.

Fekete, S.; Ganzler, K.; Fekete, J. J. Pharm. Biomed. Anal. 2011, 54, 482-

(28)

Gritti, .; Guiochon, G. J. Chromatogr., A 2012, 1221, 2-40.

(29) Ruta, J.; Guillarme, D.; Rudaz, S.; Veuthey, J.-L. J. Sep. Sci. 2010, 33, 2465-2477. (30)

Dioszegi, T. A.; Raynie, D. E. J. Chromatogr., A 2012, 1261, 107-112.

(31)

Shaaban, H.; Gorecki, T. Anal. Methods 2012, 4, 2735-2743.

(32)

Gritti, F.; Guiochon, G. J. Chromatogr., A 2011, 1218, 907-921.

(33)


(34)


(35)

Lesellier, E. J. Chromatogr., A 2012, 1228, 89-98.

(36) Tang, C.; Tsao, P.; Lin, C.; Fang, L.; Lee, S.; Wang, W. Anal. Bioanal. Chem. DOI:10.1007/s00216-012-6414-8. (37) 285.

D'Orazio, G.; Rocchi, S.; Fanali, S. J. Chromatogr., A 2012, 1255, 277-

(38)

Manns, D. C.; Mansfield, A. K. J. Chromatogr., A 2012, 1251, 111-121.

(39) Fanali, C.; Rocco, A.; Aturki, Z.; Mondello, L.; Fanali, S. J. Chromatogr., A 2012, 1234, 38-44. (40) Gomez-Caravaca, A. M.; Segura-Carretero, A.; Fernandez-Gutierrez, A.; Caboni, M. F. J. Agric. Food Chem. 2011, 59, 10815-10825.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(41) Rostagno, M. A.; Manchon, N.; D'Arrigo, M.; Guillamon, E.; Villares, A.; Garcia-Lafuente, A.; Ramos, A.; Martinez, J. A. Anal. Chim. Acta 2011, 685, 204-211. (42) Gallart-Ayala, H.; Moyano, E.; Galceran, M. T. From J. Chromatogr., A 2011, 1218, 1603-1610. (43) Gallart-Ayala, H.; Moyano, E.; Galceran, M. T. From Anal. Chim. Acta 2011, 683, 227-233. (44)

Chirita, R.; Finaru, A.; Elfakir, C. J. Chromatogr., B 2011, 879, 633-640.

(45)

DeGrasse, S. L.; DeGrasse, J. A.; Reuter, K. Toxicon 2011, 57, 179-182.

(46) Gupta, P. K.; Brown, J.; Biju, P. G.; Thaden, J.; Deutz, N. E.; Kumar, S.; Hauer-Jensen, M.; Hendrickson, H. P. Anal. Methods 2011, 3, 1759-1768. (47) Zhang, Y.; Tingley, F. D., III; Tseng, E.; Tella, M.; Yang, X.; Groeber, E.; Liu, J.; Li, W.; Schmidt, C. J.; Steenwyk, R. J. Chromatogr., B 2011, 879, 20232033. (48) Martinez-Villalba, A.; Moyano, E.; Galceran, M. T. J. Chromatogr., A 2010, 1217, 5802-5807. (49) Chauve, B.; Guillarme, D.; Cleon, P.; Veuthey, J.-L. J. Sep. Sci. 2010, 33, 752-764. (50)

Kawano, S. Rapid Commun. Mass Spectrom. 2009, 23 907-914.

(51) Cacciola, F.; Donato, P.; Giuffrida, D.; Torre, G.; Dugo, P.; Mondello, L. J. Chromatogr., A 2012, 1255, 244-251. (52) Jandera, P.; Hajek, T.; Cesla, P. J. Chromatogr., A 2011, 1218, 19952006. (53)

Jandera, P.; Hajek, T.; Cesla, P. J. Sep. Sci. 2010, 33, 1382-1397.

(54) Chocholous, P.; Kosarova, L.; Satinsky, D.; Sklenarova, H.; Solich, P. Talanta 2011, 85, 1129-1134. (55)

Guiochon, G. J. Chromatogr., A 2007, 1168, 101–168.

(56)

Svec, F. J. Chromatogr., A 2012, 1228, 250-262.

(57) Nunez, O.; Gallart-Ayala, H.; Martins, C. P. B.; Lucci, P. J. Chromatogr., A 2012, 1228, 298-323. (58) van de Meent, M. H. M.; de Jong, G. J. TrAC, Trends Anal. Chem. 2011, 30, 1809-1818. (59)

Bunch, D. R.; Wang, S. J. Sep. Sci. 2011, 34, 2003-2012.


Page 30 of 41

Page 31 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(60) Namera, A.; Miyazaki, S.; Saito, T.; Nakamoto, A. Anal. Methods 2011, 3, 2189-2200. (61) Unger, K. K.; Tanaka, N.; Machtejevas, E. Eds., Monolithic Silicas in Separation Science; Wiley-VCH Verlag GMBH, Weinheim, Germany, 2011. (62)


(63)

Gritti, F. Am. Pharm. Rev. 2012, 15, 70-74.

(64)

Cabrera, K. LCGC North Am. 2012 (Suppl.), 56-60.

(65) Trojer, L.; Greiderer, A.; Bisjak, C. P.; Wieder, W.; Heigl, N.; Huck, C. W.; Bonn, G. K. In Handbook of HPLC, 2nd Ed.; Corradini, D., Ed.; CRC Press: Boca Raton, 2010; pp 3-45. (66)

Ghanem, A.; Ikegami, T. J. Sep. Sci. 2011, 34, 1945-1957.

(67)

El Deeb, S. Chromatographia 2011, 74, 681-691.

(68)

Gawel, B.; Gawel, K.; Oeye, G. Materials 2010, 3, 2815-2833.

(69)

Walsh, Z.; Paull, B.; Macka, M. Anal. Chim. Acta 2012, 750, 28-47.

(70) Nischang, I.; Brueggemann, O.; Svec, F. Anal. Bioanal. Chem. 2010, 397, 953-960. (71) Nischang, I.; Teasdale, I.; Brueggemann, O. Anal. Bioanal. Chem. 2011, 400, 2289-2304. (72) Li, Y.; Aggarwal, P.; Tolley, H. D.; Lee, M. L. In Advances in Chromatography, Volume 50; Grushka, E.; Grinburg, N., Eds.; CRC Press: Boca Raton, 2012; pp 237-280. (73) 14.

Aggarwal, P.; Tolley, H. D.; Lee, M. L. J. Chromatogr., A 2012, 1219, 1-

(74) Li, Y.; Xie, X.; Lee, M. L.; Chen, J. J. Chromatogr., A 2011, 1218, 86088616. (75)

Chen, X.; Tolley, H. D.; Lee, M. L. J. Sep. Sci. 2011, 34, 2088-2096.

(76)

Chen, X.; Tolley, H. D.; Lee, M. L. J. Sep. Sci. 2011, 34, 2063-2071.

(77) Chen, X.; Tolley, H. D.; Lee, M. L. J. Chromatogr., A 2010, 1217, 38443854. (78)

Zhou, Q.; Zhang, P.; Jia, L. J. Sep. Sci. 2011, 34, 3303-3309.

(79) Badaloni, E.; Barbarino, M.; Cabri, W.; D'Acquarica, I.; Forte, M.; Gasparrini, F.; Giorgi, F.; Pierini, M.; Simone, P.; Ursini, O.; Villani, C. J. Chromatogr., A 2011, 1218, 3862-3875.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(80) Krajnc, N. L.; Smrekar, F.; Frankovic, V.; Strancar, A.; Podgornik, A. In Macroporous Polymers; Mattiasson, B.; Kumar, A.; Galaev, I. Y., Eds.; CRC Press: Boca Raton, 2010; pp 291-334. (81)

Zhu, T.; Row, K. H. J. Sep. Sci. 2012, 35, 1294-1302.

(82) Vujcic, S.; Ferrer, I. M.; Rivera, J. G.; Li, L.; Colon, L. A. ACS Symposium Series (2011), 1062(Interfaces and Interphases in Analytical Chemistry), 123-139. (83)

Wu, M.; Wu, R.; Zhang, Z.; Zou, H. Electrophoresis 2011, 32, 105-115.

(84) Wang, X.; Zheng, Y.; Zhang, C.; Yang, Y.; Lin, X.; Huang, G.; Xie, Z. J. Chromatogr., A 2012, 1239, 56-63. (85) Jandera, P.; Urban, J.; Skerikova, V.; Langmaier, P.; Kubickova, R.; Planeta, J. J. Chromatogr., A 2010, 1217, 22-33. (86) Zhang, Z.; Wu, M.; Wu, R.; Dong, J.; Ou, J.; Zou, H. Anal. Chem. 2011, 83, 3616-3622. (87) Arrua, R. D.; Talebi, M.; Causon, T. J.; Hilder, E. F. Anal. Chim. Acta 2012, 738, 1-12. (88)

Krenkova, J.; Foret, F.; Svec, F. J. Sep. Sci. 2012, 35, 1266-1283.

(89) Chambers, S. D.; Holcombe, T. W.; Svec, F.; Frechet, J. M. J. Anal. Chem. 2011, 83, 9478-9484. (90) Chambers, S D.; Svec, F; Frechet, J M. J. J. Chromatogr., A 2011, 1218, 2546-2552. (91) Aqel, A.; Yusuf, K.; Al-Othman, Z. A.; Badjah-Hadj-Ahmed, A. Y.; Alwarthan, A. A. Analyst (Cambridge, U. K.) 2012, 137, 4309-4317. (92) Tong, S.; Liu, Q.; Li, Y.; Zhou, W.; Jia, Q.; Duan, T. J. Chromatogr., A 2012, 1253, 22-31. (93) Tan, J.; Jiang, Z..; Li, Rong; Yan, X. TrAC, Trends Anal. Chem. 2012, 39, 207-217. (94)

Zheng, C.; Huang, Y.; Liu, Z. J. Sep. Sci. 2011, 34, 1988-2002.

(95)

He, J.; Fang, G.; Yao, Y.; Wang, S. J. Sep. Sci. 2010, 33, 3263-3271.

(96) 595.

Chen, X.; Yang, W.; Zhou, Y.; Jiao, F. J. Porous Mater. 2012, 19, 587-

(97) Mu, L.; Wang, X.; Zhao, L.; Huang, Y.; Liu, Z. J. Chromatogr., A 2011, 1218, 9236-9243.


Page 32 of 41

Page 33 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(98) Zhao, L.; Ban, L.; Zhang, Q.; Huang, Y.; Liu, Z. J. Chromatogr., A 2011, 1218, 9071-9079. (99) Calleri, E.; Ambrosini, S.; Temporini, C.; Massolini, G. J. Pharm. Biomed. Anal. 2012, 69, 64-76. (100) Brekkan, E.; Lundqvist, A.; Lundahl, P.; Biochemistry 1996, 35, 12141– 12145. (101) Gottschalk, I.; Lagerquist, C.; Zuo, S.S.; Lundqvist, A.; Lundahl, P. J. Chromatogr., B 2002, 768, 31–40. (102)

Moaddel, R.; Wainer, I.W.; Nat. Protoc. 2009, 4. 197–205.

(103)

Tetala, K. K. R.; van Beek, T. A. J. Sep. Sci. 2010, 33, 422-438.

(104)

Arrua, R. D.; Alvarez Igarzabal, C. I. J. Sep. Sci. 2011, 34, 1974-1987.

(105)

Spross, J.; Sinz, A. J. Sep. Sci. 2011, 34, 1958-1973.

(106) Pfaunmiller, E. L.; Hartmann, M.; Dupper, C. M.; Soman, S.; Hage, D. S. J. Chromatogr., A, 2012, http://dx.doi.org/10.1016/j.chroma.2012.09.009. (107) Wang, S.; Wang, M.; Su, X.; Yuan, B.; Feng, Y. Anal. Chem. 2012, 84, 7763-7770. (108) Zhang, L.; Wang, H.; Liang, Z.; Yang, K.; Zhang, L.; Zhang, Y. J. Sep. Sci. 2011, 34, 2122-2130. (109) Hou, C.; Ma, J.; Tao, D.; Shan, Y.; Liang, Z.; Zhang, L.; Zhang, Y. J. Proteome Res. 2010, 9, 4093-4101. (110) Reichelt, S.; Elsner, C.; Prager, A.; Naumov, S.; Kuballa, J.; Buchmeiser, M. R. Analyst (Cambridge, U. K.) 2012, 137, 2600-2607. (111) Shin, M. J.; Tan, L.; Jeong, M. H.; Kim, J.; Choe, W. J. Chromatogr., A 2011, 1218, 5273-5278. (112) Feng, S.; Pan, C.; Jiang, X.; Xu, S.; Zhou, H.; Ye, M.; Zou, H. Proteomics 2007, 7, 351-360. (113) Morisaka, H.; Kirino, A.; Kobayashi, K.; Ueda, M. Biosci., Biotechnol., Biochem. 2012, 76, 585-588. (114) Chen, X.; Cao, Y.; Lv, D.; Zhu, Z.; Zhang, J.; Chai, Y. J. Chromatogr., A 2012, 1242, 67-74. (115)

Rozenbrand, J.; van Bennekom, W. P. J. Sep. Sci. 2011, 34, 1934-1944.

(116) Iwasaki, M.; Sugiyama, N.; Tanaka, N.; Ishihama, Y. J. Chromatogr., A 2012, 1228, 292-297



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(117) Vitorino, R.; Krenkova, J.; Foret, F.; Domingues, P.; Amado, F. Methods Mol. Biol. (N. Y.) 2011, 790, 31-46. (118) Zhang, Z.; Wang, F.; Xu, B.; Qin, H.; Ye, M.; Zou, H. J. Chromatogr., A 2012, 1256, 136-143. (119) Kortz, L.; Helmschrodt, C.; Ceglarek, U. Anal. Bioanal. Chem. 2011, 399, 2635-2644. (120)

Zheng, Y.; Wang, X.; Ji, Y. Talanta 2012, 91, 7-17.

(121)

Wistuba, D. J. Chromatogr., A 2010, 1217, 941-952.

(122)

Chankvetadze, B. J. Sep. Sci. 2010, 33, 305-314.

(123) Zhang, Zhenbin; Wu, Ren'an; Wu, Minghuo; Zou, Hanfa Electrophoresis 2010, 31, 1457-1466. (124)

Li, H; Liu, Z TrAC, Trends Anal. Chem. 2012, 37, 148-161.

(125) Lin, Z. A.; Pang, J. L.; Yang, H.;H.; Cai, Z.;W.; Zhang, L.; Chen, G.N., 2011 47, 9675. (126) Lin, Z. A.; Pang, J. L.; Lin, Y.; Huang, H.; Cai, Z.W.; Zhang, L.; Chen. G.N. Analyst (Cambridge, U. K.), 2011, 136, 3281. (127) Yang, F.; Lin, Z.,A.; He, X.,W.; Chen, L.,X.; Zhang, Y.,K. J. Chromatogr., A 2011, 1218, 9194. (128)

Li, H. Y.; Liu, Y. C.; Liu, J.; Liu, Z. Chem. Commun. 2011, 47, 8169.

(129)

Liu, Y. C.; Ren, L.;B.; Liu, Z. Chem. Commun., 2011, 47, 5067.

(130) Li, Q.; Lue, C.; Li, H.; Liu, Y.; Wang, H.; Wang, X.; Liu, Z. J. Chromatogr., A 2012, 1256, 114-120. (131)

Liu, Y.; Lu, Y.; Liu, Z. Chem. Sci. 2012, 3, 1467-1471.

(132)

Fernandez, M.; Forteza, R.; Cerda, V. Instrum. Sci. Tech. 2012, 40, 90-99.

(133) Koblova, P.; Sklenarova, H.; Chocholous, P.; Polasek, M.; Solich, P. Talanta 2011, 84, 1273-1277. (134)

Jiang, Z.; Smith, N. W.; Liu, Z. J. Chromatogr., A 2011, 1218, 2350-2361.

(135)

Gunasena, D. N.; El Rassi, Z. Electrophoresis 2012, 33, 251-261.

(136) Nordborg, A.; Hilder, E. F.; Haddad, P. R. Annu. Rev. Anal. Chem. 2011, 4, 197-226. (137) Yang, G.; Yan, C.; Bai, L.; Li, J.; Duan, Y. Anal. Methods 2012, 4, 10981104.


Page 34 of 41

Page 35 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(138)

Yang, G.; Bai, L.; Yan, C.; Gu, Y.; Ma, J. Talanta 2011, 85, 2666-2672.

(139)

Lv, Y.; Lin, Z.; Svec, F. Anal. Chem. 2012, 84, 8457-8460.

(140) Lv, Y.; Lin, Z.; Svec, F. Analyst (Cambridge, U. K.) 2012, 137, 41144118. (141) Guerrouache, M.; Pantazaki, A.; Millot, M.; Carbonnier, B. J. Sep. Sci. 2010, 33, 787-792. (142) Silva, R. G. C.; Bottoli, C. B. G.; Collins, C. H. J. Chromatogr. Sci. 2012, 50, 649-657. (143) Lin, Z.; Huang, H.; Sun, X.; Lin, Y.; Zhang, L.; Chen, G. J. Chromatogr., A 2012, 1246, 90-97. (144) Chen, M.; Li, L.; Yuan, B.; Ma, Q.; Feng, Y. J. Chromatogr., A 2012, 1230, 54-60, (145) 192.

Chen, M.; Wei, S.; Yuan, B.; Feng, Y. J. Chromatogr., A 2012, 1228, 183-

(146)

Skerikova, V.; Jandera, P. J. Chromatogr., A 2010, 1217, 7981-7989.

(147) Lin, H.; Ou, J.; Zhang, Z.; Dong, J.; Wu, M.; Zou, H. Anal. Chem. 2012, 84, 2721-2728. (148) Gama, M. R.; da Costa Silva, R. G.; Collins, C. H.; Bottoli, C. B. G. TrAC, Trends Anal. Chem. 2012, 37, 48-60. (149) Bernal, J.; Ares, A. M.; Pol, J.; Wiedmer, S. K. J. Chromatogr., A 2011, 1218, 7438-7452. (150)

Buszewski, B.; Noga, S. Anal. Bioanal. Chem. 2012, 402, 231-247.

(151)

Guo, Y.; Gaiki, S. J. Chromatogr., A 2011, 1218, 5920-5938.

(152) Easter, R. N.; Limbach, P. A. In Handbook of Analysis of Oligonucleotides and Related Products; Bonilla, J. V.; Srivatsa, G. S., Eds.; CRC Press: Boca Raton, 2011; pp 425-438. (153) Hydrophilic Interaction Liquid Chromatography (HILIC) and Advanced Applications; Wang, P. G.; He, W., Eds. CRC Press: Boca Raton, 2011. (154)

Jandera, P. Anal. Chim. Acta 2011, 692, 1-25.

(155) Gika, H.; Theodoridis, G.; Mattivi, F.; Vrhovsek, U.; Pappa-Louisi, A. Anal. Bioanal. Chem. 2012, 404, 701-709. (156) Creek, D. J.; Jankevics, A.; Breitling, R.; Watson, D. G.; Barrett, M. P.; Burgess, K. E. V. Anal. Chem. 2011, 83, 8703-8710.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(157) Qiu, H.; Liang, X.; Sun, M.; Jiang, S. Anal. Bioanal. Chem. 2011, 399, 3307-3322. (158) Ray, S.; Takafuji, M.; Ihara, H. Analyst (Cambridge, U. K.) 2012, 137, 4907-4909. (159) Li, Y.; Xu, Z.; Feng, Y.; Liu, X.; Chen, T.; Zhang, H. Chromatographia 2011, 74, 523-530. (160) Li, Y.; Li, J.; Chen, T.; Liu, X.; Zhang, H. J. Chromatogr., A 2011, 1218, 1503-1508. (161) Qiu, H.; Loukotkova, L.; Sun, P.; Tesarova, E.; Bosakova, Z.; Armstrong, D. W. J. Chromatogr., A 2011, 1218, 270-279. (162)

Padivitage, N. L. T.; Armstrong, D. W. J. Sep. Sci. 2011, 34, 1636-1647.

(163) Kozlik, P.; Simova, V.; Kalikova, K.; Bosakova, Z.; Armstrong, D. W.; Tesarova, E. J. Chromatogr., A 2012, 1257, 58-65. (164) Qiu, H.; Wanigasekara, E.; Zhang, Y.; Tran, T.; Armstrong, D. W. J. Chromatogr., A 2011, 1218, 8075-8082. (165) Li, Y.; Feng, Y.; Chen, T.; Zhang, H. J. Chromatogr., A 2011, 1218, 59875994. (166) Guo, H.; Liu, R.; Yang, J.; Yang, B.; Liang, X.; Chu, C. J. Chromatogr., A 2012, 1223, 47-52. (167) Kotoni, D.; D'Acquarica, I.; Ciogli, A.; Villani, C.; Capitani, D.; Gasparrini, F. J. Chromatogr., A 2012, 1232, 196-211. (168)

Schuster, G.; Lindner, W. Anal. Bioanal. Chem. 2011, 400, 2539-2554.

(169)

Dai, X.; He, Y.; Wei, Y.; Gong, B. J. Sep. Sci. 2011, 34, 3115-3122.

(170) Huang, H.; Guo, H.; Xue, M.; Liu, Y.; Yang, J.; Liang, X.; Chu, C. Talanta 2011, 85, 1642-1647. (171) Li, Y.; Xu, L.; Chen, T.; Liu, X.; Xu, Z.; Zhang, H. Anal. Chim. Acta 2012, 726, 102-108. (172) 247.

Kalafut, P.; Kucera, R.; Klimes, J. J. Chromatogr., A 2012, 1232, 242-

(173) Kawachi, Y.; Ikegami, T.; Takubo, H.; Ikegami, Y.; Miyamoto, M.; Tanaka, N. J. Chromatogr., A 2011, 1218, 5903-5919. (174)

Berger, T. A. J. Chromatogr., A 2011, 1218, 4559-4568.


Page 36 of 41

Page 37 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(175) de la Puente, M. L.; Soto-Yarritu, P. Lopez; Anta, C. J. Chromatogr., A 2012, 1250, 172-181. (176)

West, C.; Khater, S.; Lesellier, E. J. Chromatogr., A 2012, 1250, 182-195.

(177) A. dos Santos Pereira, A. Giron, E. Admasu, P. Sandra, J. Sep. Sci. 2010, 33, 834. (178) Treadway, J. W.; Philibert, G. S.; Olesik, S. V. J. Chromatogr., A 2011, 1218, 5897-5902. (179)

Philibert. G. S.; Olesik, S. V. J. Chromatogr., A, 2011, 1218, 8222– 8230.

(180)

Taylor, L. T. J. Chromatogr., A 2012, 1250, 196-204.

(181)

Ashraf-Khorassani, M.; Taylor, L. T. J. Sep. Sci. 2010, 33, 1682.

(182) Patel, M. A.; Riley, F.; Ashraf-Khorassani, M.; Taylor, L. T. J. Chromatogr., A 2012, 1233, 85-90. (183)

Alpert, A. J. J. Chromatogr., A 1990, 499, 177-196.

(184)

Chester, T. L. J. Chromatogr., A, 2012, 1261, 69-77.

(185) Schiesel, S.; Laemmerhofer, M.; Lindner, W. J. Chromatogr., A 2012, 1259, 100-110. (186) Kubica, P.; Kot-Wasik, A.; Wasik, A.; Namiesnik, J.; Landowski, P. J. Chromatogr., B 2012, 907, 34-40. (187) Remoroza, C.; Cord-Landwehr, S.; Leijdekkers, A. G. M.; Moerschbacher, B. M.; Schols, H. A.; Gruppen, H. Carbohydr. Polym. 2012, 90, 41-48. (188) Lane, J. A.; Marino, K.; Rudd, P. M.; Carrington, S. D.; Slattery, H.; Hickey, R. M. J. Microbiol. Methods 2012, 90, 53-59. (189) Hernandez-Hernandez, O.; Calvillo, I.; Lebron-Aguilar, R.; Moreno, F. J.; Sanz, M. L. J. Chromatogr., A 2012, 1220, 57-67. (190) Ricochon, G.; Paris, C.; Girardin, M.; Muniglia, L. J.Chromatogr., B 2011, 879, 1529-1536. (191) Melmer, M.; Stangler, T.; Premstaller, A.; Lindner, W. J. Chromatogr., A 2011, 1218, 118-123. (192) Li, L.; Zhang, F.; Zaia, J.; Linhardt, R. J. Anal. Chem. 2012, 84, 88228829. (193) Walker, S. H.; Carlisle, B. C.; Muddiman, D. C. Anal. Chem. 2012, 84, 8198-8206.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(194) Novakova, L.; Gottvald, T.; Vlckova, H.; Trejtnar, F.; Mandikova, J.; Solich, P. J. Chromatogr., A 2012, 1259, 237-243. (195) Chen, Y.; Bicker, W.; Wu, J.; Xie, M.; Lindner, W. J. Agric. Food Chem. 2012, 60, 4243-4252. (196) Rodriguez-Gonzalo, E.; Garcia-Gomez, D.; Carabias-Martinez, R. J. Chromatogr., A 2011, 1218, 9055-9063. (197) Zhao, H.; Chen, J.; Shi, Q.; Li, X.; Zhou, W.; Zhang, D.; Zheng, L.; Cao, W.; Wang, X.; Lee, F. S. J. Sep. Sci. 2011, 34, 2594-2601. (198) Chen, P.; Li, W.; Li, Q.; Wang, Y.; Li, Z.; Ni, Y.; Koike, K. Talanta 2011, 85, 1634-1641. (199) Lin, L.; Huang, Z.; Gao, Y.; Chen, Y.; Hang, W.; Xing, J.; Yan, X. Proteomics 2012, 12, 2238-2246. (200) Sterz, K.; Scherer, G.; Krumsiek, J.; Theis, F. J.; Ecker, J. Chem. Res. Toxicol. 2012, 25, 1565-1567. (201) Spagou, K.; Wilson, I. D.; Masson, P.; Theodoridis, G.; Raikos, N.; Coen, M.; Holmes, E.; Lindon, J. C.; Plumb, R. S.; Nicholson, J. K.; Want, E. J. Anal. Chem. 2011, 83, 382-390. (202) Gika, H. G.; Theodoridis, G. A.; Vrhovsek, U.; Mattivi, F. J. Chromatogr., A 2012, 1259, 121-127. (203) Boudra, H.; Doreau, M.; Noziere, P.; Pujos-Guillot, E.; Morgavi, D. P. J. Chromatogr., A 2012, 1256, 169-176. (204) Vo Duy, S.; Besteiro, S.; Berry, L.; Perigaud, C.; Bressolle, F.; Vial, H. J.; Lefebvre-Tournier, I. Anal. Chim. Acta 2012, 739, 47-55. (205) Paglia, G.; Magnusdottir, M.; Thorlacius, S.; Sigurjonsson, O. E.; Guethmundsson, S.; Palsson, B. O.; Thiele, I. J. Chromatogr., B 2012, 898, 111120. (206) Xu, Y.; Yang, L.; Yang, F.; Xiong, Y.; Wang, Z.; Hu, Z. Metabolomics 2012, 8, 475-483. (207)

Rappold, B. A.; Grant, R. P. J. Sep. Sci. 2011, 34, 3527-3537.

(208) Li, J.; von Pfoestl, V.; Zaldivar, D.; Zhang, X.; Logothetis, N.; Rauch, A. Anal. Bioanal. Chem. 2012, 402, 2545-2554. (209) Garcia-Canaveras, J. C.; Donato, M. T.; Castell, J. V.; Lahoz, A. J. Proteome Res. 2011, 10, 4825-4834. (210) Fontanals, N.; Marce, R. M.; Borrull, F. J. Chromatogr., A 2011, 1218, 5975-5980.


Page 38 of 41

Page 39 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(211) Pasakova, I.; Gladziszova, M.; Charvatova, J.; Stariat, J.; Klimes, J.; Kovarikova, P. J. Sep. Sci. 2011, 34, 1357-1365. (212) Yoshioka, N.; Asano, M.; Kuse, A.; Mitsuhashi, T.; Nagasaki, Y.; Ueno, Y. J. Chromatogr., A 2011, 1218, 3675-3680. (213) MacIntyre, L.; Zheng, L.; Scullion, P.; Keating, P.; Watson, D. G. Metabolomics 2011, 7, 54-70. (214) Lin, L.; Huang, Z.; Gao, Y.; Yan, X.; Xing, J.; Hang, W. J. Proteome Res. 2011, 10, 1396-1405. (215) Kovarikova, P.; Stariat, J.; Klimes, J.; Hruskova, K.; Vavrova, K. J. Chromatogr., A 2011, 1218, 416-426. (216) Di Palma, S.; Boersema, P. J.; Heck, Albert J. R.; Mohammed, Shabaz Anal. Chem. 2011, 83, 3440-3447. (217) Di Palma, S.; Stange, D.; van de Wetering, M.; Clevers, H.; Heck, A. J. R.; Mohammed, S. J. Proteome Res. 2011, 10, 3814-3819. (218) Zhao, Y.; Kong, R. P. W.; Li, G.; Lam, M. P. Y.; Law, C. H.; Lee, S. M. Y.; Lam, H. C.; Chu, I. K. J. Sep. Sci. 2012, 35, 1755-1763. (219) Garbis, S. D.; Roumeliotis, T. I.; Tyritzis, S. I.; Zorpas, K. M.; Pavlakis, K.; Constantinides, C. A. Anal. Chem. 2011, 83, 708-718. (220) Zarei, M.; Sprenger, A.; Metzger, F.; Gretzmeier, C.; Dengjel, J. J. Proteome Res. 2011, 10, 3474-3483. (221) Engholm-Keller, K.; Hansen, T. A.; Palmisano, G.; Larsen, M. R. J. Proteome Res. 2011, 10, 5383-5397. (222) Kaji, H.; Shikanai, T.; Sasaki-Sawa, A.; Wen, H.; Fujita, M.; Suzuki, Y.; Sugahara, D.; Sawaki, H.; Yamauchi, Y.; Shinkawa, T.; Taoka, M.; Takahashi, N.; Isobe, T.; Narimatsu, H. J. Proteome Res. 2012, 11, 4553-4566. (223)

Wu, C.; Chen, Y.; Tai, J.; Chen, S. J. Proteome Res. 2011, 10, 1088-1097.

(224)

Nováková. L.; Vlčková, H.; Petr, S. Talanta 2012, 93, 99–105.

(225) Alpert, A. J.; Petritis, K.; Kangas, L.; Smith, R. D.; Mechtler, K.; Mitulovic, G.; Mohammed, S.; Heck, A. J. R. Anal. Chem. 2010, 82, 5253–5259 (226)

Yang, Y.; Geng, X. J. Chromatogr., A 2011, 1218, 8813-8825.

(227) Calinescu, O.; Badea, I. A.; Vladescu, L.; Meltzer, V.; Pincu, E. J. Chromatogr. Sci. 2012, 50, 335–342. (228) Kaliszan, R.; Wiczling, P. TrAC, Trends Anal. Chem. 2011, 30, 13721381.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(229) Laemmerhofer, M.; Nogueira, R.; Lindner, W. Anal. Bioanal. Chem. 2011, 400, 2517-2530. (230)

Zhang, Y.; Carr, P. W. J. Chromatogr., A 2011, 1218, 763-777.

(231) Bicker, W.; Wu, J. Y.; Yeman, H.; Albert, K.; Lindner, W. J. Chromatogr., A 2011, 1218, 882-895. (232) Zhao, W.; Wang, W.; Chang, H.; Cui, S.; Hu, K.; He, L.; Lu, K.; Liu, J.; Wu, Y.; Qian, J.; Zhang, S. J. Chromatogr., A 2012, 1251, 74-81. (233) Borges, E. M.; Euerby, M. R.; Collins, C. H. Anal. Bioanal. Chem. 2012, 402, 2043-2055. (234) Abbood, A.; Herrenknecht, C.; Proczek, G.; Descroix, S.; Rodrigo, J.; Taverna, M.; Smadja, C. Anal. Bioanal. Chem. 2011, 400, 459-468. (235)

Zhang, P.; Chen, J.; Jia, L. J. Chromatogr., A 2011, 1218, 3459-3465.

(236) Liu, X.; Pohl, C. A. In Hydrophilic Interaction Liquid Chromatography (HILIC) and Advanced Applications); Wang, P. G.; He, W., Eds. CRC Press: Boca Raton, 2011; pp 47-75. (237)

Gezici, O.; Kara, H. Talanta 2011, 85, 1472-1482.

(238) Dong, J.; Yu, L.; Zhang, X.; Xue, X.; Guo, Z.; Liang, X. Anal. Bioanal. Chem. 2011, 399, 3415-3421. (239) Wang, C.; Guo, Z.; Zhang, J.; Zeng, J.; Zhang, X.; Liang, X. J. Sep. Sci. 2011, 34, 53-58.


Page 40 of 41

Page 41 of 41

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


For TOC only


Recent Developments in High-Performance Liquid Chromatography

Recommend Documents