Capillary Electrochromatography

J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY - - v o l u m e 62 capillary e l e c tro ch ro rn a to g ra p h y...

Author: F. Svec | Z. Deyl

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J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY - - v o l u m e 62

capillary e l e c tro ch ro rn a to g ra p h y

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J O U R N A L OF C H R O M A T O G R A P H Y LIBRARY -- v o l u m e 62

capillary electroch romato graph y edited by

ZdenOk Deyl Institute of Physiology, Academy of Sciences of the Czech Republic, Prague, Czech Republic

and

v

Fran ti#ek Svec Department of Chemistry, University of California, Berkeley, USA

2001

ELSEVIER Amsterdam

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London

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New

York

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Oxford

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Shannon

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E L S E V I E R S C I E N C E B.V. S a r a B u r g e r h a r t s t r a a t 25 P.O. B o x 211, 1000 A E A m s t e r d a m , T h e N e t h e r l a n d s

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F i r s t e d i t i o n 2001 L i b r a r y o f C o n g r e s s C a t a l o g i n g in P u b l i c a t i o n D a t a C a p i l l a r y e l e c t r o c h r o m a t o g r a p h y / e d i t e d b y Z d e n e k D e y l a n d F r a n t i s e k Svec. p. c m . - - ( J o u r n a l o f c h r o m a t o g r a p h y l i b r a r y ; v. 62) I n c l u d e s b i b l i o g r a p h i c a l r e f e r e n c e s a n d index. ISBN 0-444-50432-X (hardcover) 1. C a p i l l a r y e l e c t r o p h o r e s i s . 2. C a p i l l a r y l i q u i d c h r o m a t o g r a p h y . I. D e y l , Z d e n e k . II. Svec, F r a n t i s e k . III. Series. Q P 5 1 9 . 9 . C 3 6 C 3 4 2001 543'.0894--dc21 2001018989

ISBN: 0 444 50432 X ISSN: 0301-4770 T h e p a p e r u s e d in this p u b l i c a t i o n m e e t s the r e q u i r e m e n t s o f A N S I / N I S O Z 3 9 . 4 8 - 1 9 9 2 ( P e r m a n e n c e o f Paper). P r i n t e d in T h e N e t h e r l a n d s .

V

Preface Capillary electrochromatography (CEC) is a rapidly emerging technique that adds a new dimension to current separation science. The major "news" in this method is that the hydrodynamic flow of the eluting liquid, which is typical of HPLC, is replaced by a flow driven by electro-endoosmosis. This increases considerably the selection of available separation mechanisms. For example, combinations of traditional processes such as reversed-phase- or ion-exchange- separations with electromigration techniques are now possible. Also, CEC is opening new horizons in the separation of non-polar compounds, and thus represents an alternative to the widely used micellar electrokinetic chromatography. As a matter of fact, different separation procedures such as "classical" chromatography, capillary zone electrophoresis, isoelectric focusing, and isotachophoresis, are now merging to create a unified concept of separation science that was envisioned quite some time ago. This wealth of techniques also enables their combination in a variety of operational modes to meet the continuously growing requirements of separating ever-smaller samples (both in mass and volume), removing interferences from complex matrixes, and of making on-line identification of entities separated from mixtures. As with many other instrumental methods, an increase in the complexity of the method often leads to a reduction in reliability unless the technical background has reached a level of maturity (although this is difficult to define). Thus, capillary electrochromatography benefits largely from the use of packed columns. However, this packed column technology, originally developed for HPLC, does not meet completely the needs of CEC, and new approaches are desirable. One of those new techniques is the creation of the column-bed in situ by the preparation of monolithic separation media. Obviously, history repeats itself. Many scientists will remember that the columns current in gas chromatography have evolved from packed columns and that coated open-tubular capillaries only came later. The ingenious introduction of micellar/microemulsion electrokinetic chromatography by Terabe extended the possibilities of electromigration techniques into the area of uncharged compounds. A combination of these approaches is seen to emerge in electrochromatography. As a result, today's open-tubular electrochromatography exploits the interactions of solutes with the modified inner surfaces of the capillaries, a phenomenon that many researchers active

VI in electrophoresis (and, in particular, those in protein and peptide chemistry) attempted to avoid for a number of years. The fact that electrochromatography has overcome its typical "childhood problems" is proved by the steadily increasing number of applications, as confirmed in the last chapter of this book. Clearly, electrochromatography provides the analyst with a new tool which understandably, and as with many other instrumental analytical methods, is not generally applicable to all separation tasks. However, the number of difficult separations that has been achieved elegantly using CEC within recent years is, in our opinion, sufficient to justify publication of a monograph on this subject. Within the following pages, we have attempted to provide the reader with the necessary theoretical background, description of the instrumentation, modes of operation and methods of detection, and an overview of the broad variety of applications. A common problem of all monographs concerned with rapidly developing fields is that they cannot include everything. For example, the number of papers related to CEC published during the preparation of this book increased by several tens. Similarly, it is almost impossible to cover all aspects of the subject. While some facets of the current CEC are reviewed to an adequate depth, some others are only mentioned briefly. Our intention was to publish quickly a book that may be less complete, rather than endlessly improving a work that would never see the printer's press. Since virtually nothing is constant in this world but is continuously developing, it is a fair assumption that CEC will also be developed further in the very near future, and a new monograph might be required. Therefore, we would appreciate greatly any comments and suggestion from readers which will help us to improve future editions.

Frantigek ~;vec Zden6k Deyl

Berkeley and Prague December 2000

VII

List of Contributors Luis A. Col6n

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA Anna Dermaux

State University of Gent, Organic Chemistry Department, Krijgslaan 281 (S. 4.), B 9000 GENT, Belgium Zden6k Deyl

Institute of Physiology, Academy of Sciences of the Czech Republic, Videhsk6 1083, CZ-14220 Praha 4, Czech Republic Melvin R. Euerby

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, United Kingdom Adam M. Fermier

The R.W. Johnson Pharmaceutical Research Institute, Science and New Technology~Analytical Development, OMP Bld. B-236, 1000 Route 202, Raritan, NJ 08869, USA Georg H61zl

Institute of Analytical Chemistry and Radiochemistry, Leopold-FranzensUniversity, Innrain 52 a, A-6020 Innsbruck, Austria Csaba Horwith

Department of Chemical Engineering, Yale University, New Haven, CT, USA Christian G. Huber

Institute of Analytical Chemistry and Radiochemistry, Leopold-FranzensUniversity, Innrain 52 a, A-6020 Innsbruck, Austria Christopher M. Johnson

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, United Kingdom Hiroshi Kobayashi

Department of Polymer Science and Engineering, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan Todd D. Maloney

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA

VIII Maria T. Matyska

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA Alan P. McKeown

AstraZeneca R&D Charnwood, Bakewell Road, Loughborough, Leicestershire, LE11 5RH, UnitedKingdom Ivan Mik~ik

Institute of Physiology, Academy of Sciences of the Czech Republic, Videhsk6 1083, CZ-14220 Praha 4, Czech Republic Joseph J. Pesek

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA Anurag S. Rathore

Bioprocess Sciences, Pharmacia Corp., Mail Code GG3K, 700 Chesterfield Parkway North, Chesterfield, MO 63198 Gerard P. Rozing

Agilent Technologies GmbH, Waldbronn Analytical Division, P.O. Box 1280, D 76337 Waldbronn Pat Sandra

State University of Gent, Organic Chemistry Department, Krijgslaan 281 (S. 4.), B 9000 GENT, Belgium Volker Schurig

Institute of Organic Chemistry, University of Tidbingen, Auf der Morgenstelle 18, 72076 Tiibingen, Germany Franti~ek ~;vec

Department of Chemistry, University of California, Berkeley, CA 94720-1460, USA Nobuo Tanaka

Department of Polymer Science and Engineering, Kyoto Institute of Technology, Matsugasaki, Sakyo-ku, Kyoto 606-8585, Japan Dorothee Wistuba

Institute of Organic Chemistry, University of Tiibingen, Auf der Morgenstelle 18, 72076 Tfibingen, Germany

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . List of Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 1

1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

V VII

Migration of Charged Sample Components and Electroosmotic Flow in Packed Capillary Columns ...................... Anurag S. Rathore and Csaba Horv~th

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F l o w o f ions in o p e n tubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E l e c t r o o s m o t i c f l o w in o p e n tubes . . . . . . . . . . . . . . . . . . . . . . . . . . F l o w o f ions in p a c k e d c o l u m n s . . . . . . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow through packed columns . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S y m b o l s and a b b r e v i a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 2

Instrumentation for Capillary Electrochromatography

........

1 2 3 5 6 12 35 36 37

39

Gerard P. Rozing, Anna Dermaux and Pat Sandra 2.1 2.2 2.3 2.4 2.5 2.6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation requirements . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gradient-CEC instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . S u m m a r y and c o n c l u s i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 3

Modes of CEC Separation . . . . . . . . . . . . . . . . . . . . . . . . .

40 41 58 75 82 83

87

Christopher M. Johnson, Alan P. McKeown and Melvin R. Euerby 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Unmodified packings . . . . . . . . . . . . . . . . . . . . . . . . Modified packings . . . . . . . . . . . . . . . . . . . . . . . . . Chiral stationary p h a s e s . . . . . . . . . . . . . . . . . . . . . . Gel C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Monoliths . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Size e x c l u s i o n C E C . . . . . . . . . . . . . . . . . . . . . . . .

3.9 3.10 3.11 3.12

Gradient CEC . . . . . . . . . . . . . . . . Selectivity c o m p a r e d with L C . . . . . . G u i d e l i n e s for the analysis o f acidic basic Conclusions . . . . . . . . . . . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

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. . . . . . . .

. . . . . . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . and neutral c o m p o u n d s . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. . . .

88 89 89 94 100 101

101 102 103 103 104 106

3.13 3.14

106 106

Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 4

Packed Bed Columns . . . . . . . . . . . . . . . . . . . . . . . . . . .

111

Luis A. Col6n, Todd D. Maloney and Adam M. Fermier 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8

Introduction . . . . . . C o l u m n fabrication . . Packing methods . . . Comparison of packing Conclusions . . . . . . Acknowledgement . . Abbreviations . . . . . References . . . . . .

Chapter 5

. . . . . . . . . . . . . . . . . . . . . procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Capillary Electrochromatography on Monolithic Silica Columns

112 112 150 156

158 158 159 159

. . 165

Nobuo Tanaka and Hiroshi Kobayashi 5.1 5.2 5.3 5.4 5.5 5.6

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . M o n o l i t h i c silica c o l u m n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preparation procedure o f silica m o n o l i t h s from silane m o n o m e r s . . . . . . . . . Structural properties o f m o n o l i t h i c silica c o l u m n s prepared in a capillary . . . . . P e r f o r m a n c e o f m o n o l i t h i c silica c o l u m n s in C E C . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 6

165 167

169 170 173

180

Capillary Column Technology: Continuous Polymer Monoliths . . . 183

Franti~ek Svec 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 6.10 6.11 6.12

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 184 R e p l a c e a b l e p o l y m e r i c stationary phases . . . . . . . . . . . . . . . . . . . . . . 185 P o l y m e r gels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 H i g h l y crosslinked a c r y l a m i d e - b a s e d m o n o l i t h s . . . . . . . . . . . . . . . . . . 189 I m p r i n t e d enantioselective m o n o l i t h s . . . . . . . . . . . . . . . . . . . . . . . 206 Polystyrene-based monoliths . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 M e t h a c r y l a t e ester-based m o n o l i t h i c c o l u m n s . . . . . . . . . . . . . . . . . . . 212 A s s e s s m e n t o f p o r o u s structure . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 Effects o f properties on the separation ability . . . . . . . . . . . . . . . . . . . 223 Other applications o f p o r o u s p o l y m e r m o n o l i t h s in C E C c o l u m n t e c h n o l o g y . . . 232 Acknowledgment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

Chapter 7

Open Tubular Approaches to Capillary Electrochromatography

. . 241

Joseph J. Pesek and Maria T. Matyska 7.1 7.2 7.3 7.4 7.5 7.6 7.7

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l etching process . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l m o d i f i c a t i o n process . . . . . . . . . . . . . . . . . . . . . . . . . . C h a r a c t e r i z a t i o n o f etched, c h e m i c a l l y m o d i f i e d capillaries . . . . . . . . . . . . Applications of O T C E C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

242 . .

. . .

244 245 247 257 266 267

XI

7.8

References

Chapter 8

8.1 8.2 8.3 8.4 8.5 8.6 8.7

Hyphenation of Capillary Electrochromatography and Mass Spectrometry: Instrumental Aspects, Separation Systems, and Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Christian G. Huber and Georg HOlzl

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I n s t r u m e n t a t i o n and t e c h n o l o g y for c o u p l i n g o f C E C and M S .......... Stationary p h a s e - m o b i l e p h a s e systems used for C E C - - M S ........... O p t i m i z a t i o n o f e l e c t r o c h r o m a t o g r a p h i c and m a s s s p e c t r o m e t r i c c o n d i t i o n s Examples of application . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Chapter 9

9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8 9.9 9.10

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

268

271

.

272 273 296 . . . 298 . 305 . 312 . 313

Pressure Supported CEC: a High-Efficiency Technique for Enantiomer Separation . . . . . . . . . . . . . . . . . . . . . . . Dorothee Wistuba and Volker Schurig

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . T e c h n i q u e s o f pressurizing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Instrumentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A d v a n t a g e s and d i s a d v a n t a g e s o f pressurized C E C . . . . . . . . . . . . . . . . C o n c l u s i o n and future trends . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

317 318 319 321 325 326 335 336 337 337 337

Chapter 10 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zden~k Deyl and Ivan Mik~/k

341

10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 10.12 10.13 10.14

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preconcentration procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ketones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Carbohydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatty acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Triglycerides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steroids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A m i n o acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . N u c l e i c acids constituents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antibiotics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pesticides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

341 342 343 347 348 350 352 355 358 359 362 363 368 369

10.15 A v a i l a b l e applications ( s u m m a r i z i n g Table) . . . . . . . . . . . . . . . . . . . . 10.16 R e f e r e n c e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

369 413

Index of Compounds Separated . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

421


Chapter 1

Migration of Charged Sample Components and Electroosmotic Flow in Packed Capillary Columns Anurag S. RATHORE* and Csaba HORV/kTH

Department of Chemical Engineering, Yale University, New Haven, CT, USA *Present address: Bioprocess Sciences, Pharmacia Corp., Mail Code GG3K, 700 Chesterfield Parkway North, Chesterfield, MO 63198

CONTENTS

1.1 1.2 1.3 1.4

1.5

1.6 1.7 1.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Flow of ions in open tubes . . . . . . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow in open tubes . . . . . . . . . . . . . . . . . . . . . . . Flow of ions in packed columns . . . . . . . . . . . . . . . . . . . . . . . . 1.4.1 Conservation of electric current . . . . . . . . . . . . . . . . . . . . 1.4.2 Evaluation of conductivity of the packed and open segments . . . . . 1.4.3 Evaluation of the potential drop across the packed and open segments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.4 Evaluation of the equivalent length of the packed segment . . . . . . 1.4.5 Evaluation of the electric field strengths in the two segments . . . . 1.4.6 Conductivity ratio in CEC . . . . . . . . . . . . . . . . . . . . . . Electroosmotic flow through packed columns . . . . . . . . . . . . . . . . 1.5.1 Overbeek's model for EOF in porous media . . . . . . . . . . . . 1.5.2 Effect of charged capillary wall . . . . . . . . . . . . . . . . . . . 1.5.3 Conservation of volumetric flow rate . . . . . . . . . . . . . . . . 1.5.4 Flow equalizing intersegmental pressure . . . . . . . . . . . . . . 1.5.5 Flow velocities in the packed and open segments . . . . . . . . . . 1.5.6 EOF inpacked columns . . . . . . . . . . . . . . . . . . . . . . . 1.5.7 Migration times, velocities and M-factors . . . . . . . . . . . . . . 1.5.8 Higher separation efficiencies in CEC . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Symbols and abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 5 6 6 6 7 8 10 11 12 12 14 18 20 23 26 27 32 35 36 37

2

Chapter 1

1.1 INTRODUCTION Capillary electrochromatography (CEC) is a liquid phase analytical separation technique that is carried out with capillary columns by electroosmotically driven mobile phase at high electric field strengths in an apparatus similar to that used in capillary zone electrophoresis (CZE). The history of CEC could be traced to almost 60 years back when Strain [1,2] applied electric field across an adsorption column to demonstrate higher selectivity due to combination of electrophoretic and chromatographic separation forces. Almost thirty years later in 1974, Pretorius et al. suggested the use of EOF as "pumping mechanism" alternative to pressure driven flow in order to expand the scope of the then newfangled technique of HPLC [3]. The viability of CEC in packed capillary columns was demonstrated by Jorgenson and Lukacs in 1981 [4] and examined in more detail by Knox and Grant in 1987 [5,6]. One of the primary reasons of interest in the technique is the mixed separation mechanism of CEC that is borrowed from both HPLC and CZE [7]. In order to exploit the potential of CEC, it is of utmost importance to successfully control and optimize electrochromatographic conditions to generate high EOF velocities. Several experimental and theoretical studies have been performed and have tried to address these issues [8-30]. However, theoretical understanding of the EOF and the associated transport phenomena in porous media such as packed columns in CEC requires an exact description of the flow field in porous media under conditions prevalent in CEC by simultaneously solving the Poisson-Boltzmann and NavierStokes equations, which remains an unsolved and complicated task [9,10,31-54]. Columns employed so far in capillary electrochromatography (CEC) contain both a packed and an open segments with concomitant changes of the electric field strength and the flow velocity at the interface of these two segments in such columns. Figure 1.1 shows the schematics of a CEC column of total length L, which consists of a

Packed Floating segment tretaining rit ~ ,t

Fixed retaining frit

:

Open Lopen se/~l

::1

I~ L ~n L ---packed ~ op~:, ~~-" o~en,2 Detection window

Fig. 1.1. Schematic illustration of a CEC column consisting of a packed and an open segment. The latter is divided into pre-detection and post-detection open segments by the detector window.

Migration and Electroosmotic Flow

3

packed and an open segment of lengths Lpackedand Lopen, respectively. This complicates the exact calculation of the pertinent CEC parameters in the packed and the open segments, the column design and the interpretation of electrochromatographic data [18,20,21,24,26,27]. In view of the great present interest in CEC, we hope that this chapter reveals the complexity of the electrokinetic phenomena underlying EOF in porous media and will stimulate the experimental and theoretical work necessary for a better understanding of the physicochemical basis of CEC and to develop this technique into a powerful analytical tool. The goal of this chapter is to review previous work, to reveal the complexity of the electrokinetic phenomena underlying electroosmotic flow (EOF) in porous media and to compare the case of an open capillary that is used in capillary zone electrophoresis (CZE). It follows from the conservation of the volumetric flow rate that in most cases a "flow-equalizing intersegmental pressure", which is different from the pressures at the two ends of the column, develops at the interface of the packed and open segments and has significant effect on the magnitude as well as the radial distribution of the flow velocity in the open segment. A framework is presented that uses measurements of current and EOF performed on CEC columns for the evaluation of parameters such as the conductivity ratio and the interstitial EOF mobility that are useful tools for characterizing them. The actual EOF mobility, that is obtained after taking into account the porosity and tortuosity of the packing, is a better measure of surface properties of the packing. Further, a modified form of Overbeek's model that was originally developed for porous media of infinite dimensions has been used to account for the wall effect in the packed capillary columns used in CEC. Finally, migration of charged components in a CEC column is simulated and expressions are presented for estimating the retention time if the chromatographic and electrophoretic properties of the sample components are known. It is shown that, by varying the length of the packed segment and maintaining same the total length of the CEC column and the position of the detection window, the balance of the chromatographic and electrophoretic forces can be shifted and the selectivity can be adjusted if the separation involves the interplay of both mechanisms. 1.2 FLOW OF IONS IN OPEN TUBES

When an open tube with fixed charges at the tube wall is filled with an electrolyte solution, the ionic atmosphere forms an electrical double layer [31-33]. Since the double layer has a higher concentration of counterions than the bulk solution, electroneutrality requires that the bulk electrolyte outside the double layer has the same amount of excessive coions. Flow of ions in an open capillary tube occurs via one of the three possible modes. References pp. 37-38

4

Chapter 1

First is ionic conduction through the bulk electrolyte, where ions migrate under the influence of the electric field by virtue of their electrophoretic (ionic) mobilities. Second is convection of the bulk electrolyte, where the mobile phase is voltage and/or pressure driven and carries the excess charge in the bulk with it [34-36]. This contribution to the total conductivity is found to be negligibly small at ionic concentrations higher than 1 mM [34,35]. Third is ionic conduction through the double layer at the tube wall due to migration of the excess counterions under the influence of the electric field [34,36-38]. The contribution of surface conduction to the total conductivity is significant only at ionic concentrations smaller than 10 mM and zeta potentials smaller than 150 mV [34,36-38]. When ionic conduction through the bulk electrolyte is the dominant mechanism of ionic migration, the conductivity of an electrolyte filled cylindrical capillary, CYopen,is expressed as [18]

i Lopen (~open : Vopen Aope n

(1.1)

where, i is the current flowing through a capillary of length, Lopen, and cross-sectional area, Aopen, when a potential drop, Vopen, is applied across it. The conductivity of an electrolyte solution is an intensive property that is independent of the length or diameter of the capillary tube and can be expressed in terms of the concentration charge and mobility of the constituent ions as follows [ 18,39]

Oopen = [ 72 ~

Z2 Vj Cj

(1.2)

J

where, F is the Faraday constant and zj, vj and cj are the valency, mobility and molar concentration of the j th ionic species, respectively. Equation 1.2 can be rewritten as

~open= F2 C Z z2 Vj Xj

(1.3)

J

where, C is the molar concentration of the buffer and xj = cj/C is the number of moles of jth ionic species per mole of buffer. For weak electrolytes, xj depends on the dissociation of the ionogenic species and thus on the pH. It follows from Eq. 1.3 that


5

the conductivity of sufficiently dilute electrolytes increases linearly with the molar concentration of the electrolyte and the slope depending on the charge, size and concentration of the ionic species. 1.3 ELECTROOSMOTIC FLOW IN OPEN TUBES

In open tubes with thin double layers and when there is no polarization, the EOF mobility, kteo, open,

can

be expressed by the following relationship [40-44]

~: eo ~.~ ~eo,open

-"

(1.4)

"

where, e is the dielectric constant of the medium, eo is the permittivity of the vacuum, and rl is the viscosity of the bulk solution. ~w is the zeta potential of the wall and is defined as potential on a hypothetical 'surface of shear' close to the tube wall that can be assumed to exist so that while the bulk electrolyte beyond this surface is moving, there is no motion between this surface and the charged wall. The mobility, as defined by Eq. 1.4, depends on the properties of the charged surface and the bulk electrolyte but is independent of the capillary diameter or the applied voltage. When the surface

(z e qJo~

charge is independent of the electrolyte concentration and exp~ kB T ) >>1 for the simpler case of z:z electrolytes [33]

Wo= (ksT~ logC + constant

(1.5)

~,ze)

where, ~o is the surface potential, kB is the Boltzmann constant, T is the absolute temperature and e is the elementary charge. Since the zeta potential is of the same order as the surface potential for the present case, the above equation can be rewritten to relate the EOF mobility to the buffer concentration as follows

~eo,open

=

eeo~_(e_eoksT)logC+constantq Tlze

Referencespp. 37-38

(1.6)

6

Chapter 1

According to Eq. 1.6, the dependence of the EOF mobility on the logarithmic electrolyte concentration is linear. In practice, the EOF mobility in the bulk electrolyte is estimated from migration data obtained with a suitable neutral and inert tracer in an open tube by using the following expression

LdL

(1.7)

~[eo,open- to,open V

where, Ld is the distance between the inlet and the point of detection of the capillary and to, open is the migration time of the tracer in an open tube of length, L, with an applied voltage, V. 1.4 FLOW OF IONS IN PACKED COLUMNS 1.4.1 Conservation of electric current Besides the three modes of ionic migration discussed above for an electrolyte filled open silica capillary, current in a packed column can also flow via conduction through the stationary phase. However, when ionic conduction through the bulk electrolyte solution is still the primary form of ionic migration, conductivity of a packed column, ~packed, can be expressed along similar lines as Eq. 1.1 in the following manner [18]

i' Lpacked

(1.8)

(~packed = VpackedAopen

where, i' is the current flowing through the intraparticulate and interstitial mobile phase spaces of a CEC column fully packed with a stationary phase, length of the column and Vpackedis the applied voltage.

Lpacked is the

1.4.2 Evaluation of conductivity of the packed and open segments Columns in CEC often have a packed and an open segment with a detection window in between. The conductivities of the open and packed segments,

(Yopenand

Crpacked, are then evaluated from the currents measured with the capillary tubing in the absence, iopen, and presence, ipacked, of the packing by using Eqs. 1.1 and 1.8 as


7

follows

(1.9)

L iopen (Yopen= V A open

and (1.10)

L iopen ipackedLpacked (Yopen-- V [iopenL - ipackedLopen]Aopen

where, Lopen and Lpacked a r e the respective lengths of the open and the packed segments of the CEC column. 1.4.3 Evaluation of the potential drop across the packed and open segments

Once CYopenand Cypackeda r e known, the potential drops across the two segments of either column with packed length, Lpacked, are evaluated by using the following expressions [ 18,20]

I

I

(1.11)

(YopenLpacked+ (YpackedLopen

Vopen--V_Vpacked:V~~--VI

(YpackedL~ I (YopenLpackedd- (YpackedLopen

(1.12)

where, Ropen, Rpacked and R are resistances of the open segment, the packed segment and the total column, respectively. Equations 1.11 and 1.12 express that the total potential drop, V, is distributed across the packed and open segments of the column according to the relative magnitude of their resistances. Figure 1.2 shows that with increasing dimensionless packed length, ~, = Lpacked/L, the total resistance of the column and hence the potential drops across the two segments vary nonlinearly according to Eqs. 1.11 and 1.12. As ~, goes from 0 to 1, the two potential drops,

Vpackedand Vopen, monotonicaly increase and decrease from zero

to V and V to zero, respectively. Further, a higher conductivity ratio, CYopen/Cypacked, means a higher resistivity of the packed segment. Thus, a higher potential drop across References pp. 37-38

8

Chapter 1

20

.

.

.

.

|

.

.

.

.

i

.

.

.

.

|

.

.

.

.

Packed segment

>~e

i,i o p e n segmen t . . . . . . ~

15

cs

2 ~ e-, o o

(Yopen /(Ypacked

\, /1.5 _ \X/3.1 10 **~

1.5

-.

5

"

~

% ~

9"'~.. -..

10

~ 9.,..

~. 9-. ......

.

0

.

.

.

.

i

0.25

.

.

.

.

|

0.5

.

.

.

.

,

0.75

.

.

.

~

.

0

0.25

0.5

0.75

1

Dimensionless packed length, ;~ Fig. 1.2. Plots of the potential drop across the packed and the open segment of a CEC column against the dimensionless packed length, ~,, with conductivity ratio, CYopen/CYpacked,as the parameter. The respective currents with columns having packed segments of 0, 10 and 20 cm were reported as 6.6, 3.9 and 2.6 pA, respectively. The conditions were: fused silica capillary, 50 p,m x 30 cm; applied voltage, 18 kV; column packing, 3.5 mm Zorbax ODS, 80 A; mobile phase, 50 % (v/v) ACN in 10 mM sodium borate, pH 8.0.

the packed segment is required to generate current equal to that in the open segment causing an increase in the nonlinearity of V vs ~ plots in Figure 1.2. Since the total potential drop is kept constant, the plots of Vpacked and Vopen against ~, are mirror images of each other. The data plotted in Figure 1.2 was obtained by measuring the current in columns differing only in the length of packed segment, Lpacked, under otherwise identical condition [ 19]. The conductivity ratio, CYopen/CYpacked,was calculated to be 3.1 and the potential drops across the two segments, Vpacked and Vopen, were evaluated using Eqs. 1.11 and 1.12. 1.4.4 Evaluation of the equivalent length of the packed segment

For the sake of simplicity, we may replace the packed segment by a hypothetical open tube of length, Le, that is the same as the distance traveled by the neutral and inert tracer in the packed segment. The lumen of this hypothetical tube, Apacked, is assumed to be the same as the free cross-sectional area of the packed column so that 2 Apacked = gae , where, ae is the radius of the hypothetical tube. The equivalent length,

Le, is determined from the ratio of the conductivities of the packed and the open


9

O'open/~packed -o

4

0 O.

3

,.J ............... .

.

3.]

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.

.

I

0

.

1.5

,

,

,

,

0.25

I

0.5

,

~

,

,

I

0.75

1

Dimensionless packed length, Fig. 1.3. Plots of the ratio of the equivalent and the actual lengths of the packed segment, Le/Lpacked, against ~ with conductivity ratio as the parameter. Conditions same as in Figure 1.2.

segments of the column, as [20,24,43]

Le L 41 +LPLked ( (Yopen ) _( LopenI Lpacked-- Lpacked ~(~packed-- 1 ~Lpacked)

(1.13)

Altematively, the equivalent length can be evaluated directly from the current measurements in the absence and presence of the packing, topen and Zpacked, respectively.

_ q ioen I Lpacked Lpacked i;acked ~LpackedJ According to Eq. 1.13, Le depends on the length of the packed segment, Lpacked, as well as the conductivities of both the mobile and stationary phases. The ratio of lengths, Le/Lpacked, as a function of ~ is plotted in Figure 1.3 with the conductivity

Referencespp. 37-38

10

Chapter 1

ratio, t~open/CYpacked,a s the parameter. A higher t~open/t~packedratio signifies an increase in the resistivity of the packed segment which is equivalent to an increase in the Le/Lpacked. Further, if the resistivities of the two segments are close, as for C~open/t~packed= 1.5 or 3.1, Le/Lpacked is nearly a constant as assumed in the following discussion. 1.4.5 Evaluation of the electric field strengths in the two segments

The actual electric field (under which the migrants travel) in the packed segment, V p a c k e d , and the equiva-

Epacked, can be evaluated from the pertinent potential drop, lent length, Le, as

Epacked =

Vp,~a Le

(1.15)

whereas, the electric field in the open segment is obtained by the relationship

Vopen

(1.16)

gopen = Zopen

Evidently, both Epacked and Eopen a r e quite different from the "fictitious electric field", E*, that is obtained by dividing the total potential drop across the column, V, by the total column length, L.

E* =__V L

(1.17)

Clear distinction between Epacked, Eopen and E is of particular importance when accurate knowledge of the electric field strength is required for evaluation of the electroosmotic and electrophoretic mobilities from the observed migration times. Similar situation arises in other systems that are subject to similar conservation principles, e.g., displacement electrophoresis. Figure 1.4 illustrates plots of the above three electric field strengths against the dimensionless packed length ~, according to Eqs. 1.15, 1.16 and 1.17 with

CYopen/t~packed=

3.1. It is seen that, the field strength is higher in the packed than in


11

TE*

I Eopoo

LU

-J~

r

100

|

K,

80

"'~,

oLU

!

|

,E packed

~'-~

L

~E

! Eop en

E packed /X

.

"-,~,

60

E

...........................

~ .............................................................

40

.m

dl=a

o

20

u.I

0

,

,

|

O

I

,

,

,

,

I

,

,

,

,

I

,

.25 0.5 0.75 Dimensionless packed length,;~

,

,

,

1

Fig. 1.4. Plots of the actual, Epackedand Eopen, and the fictitious, E*, electric field strengths in the packed and the open segments against )~ for Case I with (yopen/CYpacked = 3.1. At the top, the discontinuity in the electric field strength at the interface of the two column segments is illustrated. Rest of the conditions as in Figure 1.2. the open segment, i.e., Epacked > Eopen. Both field strengths, Epacked and Eopen, increase with decreasing )~ and concomitantly the mobile phase velocity increases in both segments. Figure 1.4 also illustrates the inequality of the actual electric fields in the two segments that serves as a caveat that instead of the fictitious electric field strength, E*, the actual electric field strengths, Epacked and Eopen, should be evaluated and reported together with other chromatographic conditions in CEC. 1.4.6 Conductivity ratio in CEC The ratio of the conductivities of the packed and the open segments of a CEC column, % is given by the following relationship

Clp~kea

(1.18)

(Y open

The reciprocal of q~ is often called "formation factor" in reservoir engineering and

Referencespp. 37-38

12

Chapter 1

geology [45-47]. It is independent of mobile phase properties except when the ionization of the liquid is affected by the stationary phase and therefore, offers a simple means to characterize properties of the stationary phase. The conductivity ratio has been used to estimate the porosity of rock samples [45,47] and is related to the electrokinetic porosity, eT, by Archie's law as follows

(Ypacked q)----- ~ - -

(1.19)

m ET

(Y open

where, m is an empirical constant. The electrokinetic porosity in Eq. 1.19 depends on the morphology of the packing particles and is not necessarily the same as the porosity employed in chromatography. When the porosity of the media is greater than 0.2, as is the case generally, m = 1.5 provides a very close approximation to the experimental data and so this value will be used here [46,47,49,50]. 1.5 ELECTROOSMOTIC FLOW THROUGH PACKED COLUMNS 1 . 5 . 1 0 v e r b e e k ' s model for EOF in porous media

The following analysis is based on Overbeek's work [43,52,53] and is valid for porous/nonporous packing particles of any arbitrary shape as depicted in Figure 1.5. The assumptions are that the particles be non-conducting, have uniform zeta potential and a double layer thin compared to the radius of the pores in the plug. Overbeek, upon integration over the whole interstitial volume of the bed, obtained the following expression for the average velocity

(1.20) " ~ U P > - - W c cl I v f fc p

dVc=- ~r~l ~

d~

where, ~ is the zeta potential at the surface, Vc and Vcfare the total column Vvolume and the volume of the interstitial space, respectively. The integration is performed over Vcf only since flow is only in the interstices. Up is electroosmotic velocity that is generated locally at the packing surface and is given by an expression similar to Smoluchowski's equation for the EOF, as follows

Migration a n d Electroosmotic Flow

13

Lpacked

/

/

Packing

Flow oath of length, Le and tortuosity, "c = L e/Lpacked

Fig. 1.5. Schematic illustration of the tortuous flow path in the packed segment of a CEC column.

Up

~

~eo~E

(1.21)

lq

For the current, i, the following relationship holds

i = OpackedE = ~(YopenIV? d

(1.22)

where, ~packedand Oopen are the conductivities of a completely packed column and an open tube, both filled with the electrolyte solution, respectively. Combination of Eqs. 1.20 and 1.22 yields the average velocity as

= e eo ~ E (Opacked~

1]

(1.23)

~ (Yopen) !

with the conductivity ratio,

Opacked/Oopen,that is readily determinable experimentally.

Equation 1.23 was derived by Overbeek and Wijga [52] and Overbeek [53] to describe EOF in a porous medium without boundaries and will be used in the next section to express the effect of the capillary wall on the flow distribution inside the column.

References pp. 3 7-38

14

Chapter 1

1.5.2 Effect of charged capillary wall It is assumed that the EOF is generated only at the charged wall and the packing particles are uncharged. The flow can then be visualized in the form of very thin annuli of liquid in the packed column. Each annulus faces a force in the forward direction (the direction of EOF) from the annulus enveloping it and a force in the backward direction from the annulus inside it. The inertia terms and the compressibility of the fluid are assumed to be negligibly small. The net viscous force, Fv, on such an annulus of unit volume in absence of any particles is given by

Fv = Ar -d-rrd(r d u~rW)d)

(1.24)

where, r is the radial coordinate and Urw is the local velocity in the axial direction. In such a column with charged tube wall and uncharged packing, the flow velocity would be a rapidly fluctuating function of radial position with zero value at the surface of the uncharged particle and maximum in the intraparticular space. Hence, the velocity under consideration, Urw, is more like a volume average velocity for the volume element. Besides the viscous forces, the fluid in the shell also experiences a drag force from the packing particles in the shell. The total drag force, Fd, offered by spherical packing particles of diameter dp that are located far enough from each other to act like isolated spheres, is given by the product of the drag force by an isolated sphere and the number of spherical particles in a shell of unit volume as follows

1-ST) Fa = (6 n 1"1dp Urw) 4 ~ d3p/3

(1.25)

where, ~;T is the total porosity of the column. Since in packed columns the particles are in close contact with each other, the actual drag force is different than that given by Eq. 1.25. This is corrected for by introducing the dimensionless packing parameter, c~, which depends on the structure of packing and shape of the particles and should be easily determinable from experimental data. Performing the balance between the viscous and the drag forces and simplifying the equations we have that

Migration and Electroosmotic Flow 1 d (r du~w~=9 c~(1-~:r) u,.~ 132

5

4

15 (1.26)

where 13 is another dimensionless constant that is readily evaluated if et and ~;T are known from the following relationship

./

(1.27)

13=3-N et (1 - ~;'' 2

The boundary conditions for solving the differential equation as given in Eq. 1.26 are

d u~ _ 0

at r = 0

(1.28)

u~=uw

atr-a

(1.29)

dr

and

where, Uw is the EOF velocity at the wall and a is the tube diameter. Equation 1.29 simply reflects no slip condition at the plane of shear that is very close to the tube wall for thin double layers. Solution of the system of Eqs. 1.26, 1.28 and 1.29 for the potential distribution can be found in the literature [39,42] and written for the local velocity as

io (~ ~/r ] u :uw Io(

(1.30)

a/r

where, Io is the Bessel's function of the zeroth order. For ~r/dp greater than 3.5, which is generally the case, the zeroth order Bessel's function can be approximated by

Referencespp. 37-38

16

Chapter 1

!

l

Packing particle

Uw

Electrosmotic

velocity

Fig. 1.6. Schematic illustration of loci of the wall effect where EOF decays in the case of charged tube wall and uncharged packing particles.

e ~r/dp Io (f3 r/dp)= 42 rt [3r/dp

(1.31

)

which in turn can be substituted back into Eq. 1.30 to give the simplified solution as [11]

I e ~'/4 {2x[3a/dp I u~ = u~ ~12 rt [3r/dp e f3a/dp

= Uw

I~f~)e~("-')/4

(1.32)

Equation 1.32 for the region close to the wall is illustrated in Fig. 1.6. The flow velocity is maximal at the plane of shear and then decays quickly as we move away from the wall. This should not be surprising as in a column packed with uncharged particles, EOF is generated at the tube wall and the interior of the tube contributes only the drag resistance. It should be noted that the flow velocity falls significantly as soon as we move a distance of one particle diameter away from the column wall. Previous experimental studies on packed beds that have been published in the literature support these findings [55].


17

The average velocity in the column for the above case can be easily determined using the velocity profile given in Eq. 1.32 and accounting for the tortuosity in the bed as

- -

~o27~rurwd a2 ~,((Ypackedl=cYopen Uwl(YopenCYpacked II.a21o(_~a/dp))2 ~orlo(~r/dp)dr(1.33)

that can be simlified for the case when [~r/dp is greater than 3.5 as follows [ 11]

I

< blzw> = blw O a'e ll (~open

a2

l

(1.34)

Equations 1.34 suggests that the average velocity in the packed column varies linearly with the dimensionless particle diameter, dp/a. Under conditions employed in CEC, the EOF is generated not only at the capillary wall but also at the surface of the packing. When the zeta potential of the wall is same as the packing (~w = ~p), the velocity profile should be flat according to Overbeek's expression in Eq. 1.23. For the case when the zeta potentials at the tube wall and the particle surface are not equal, the total velocity could be evaluated by adding a term to Overbeek's velocity expression to account for the wall effect. This term would be given for the local and average velocities with the zeta potential of the wall being replaced by the 'excess zeta potential' on the wall (~w - ~p) that is responsible for the wall effect, i.e.,

(1.3:5)

where, Ur is the net local velocity from both contributions. Figure 1.7 illustrates the effect of the magnitude of the excess zeta potential on the radial profile of the EOF velocity. The plots show that the wall effect is limited to a narrow annulus at the wall that increases in width with the magnitude of the excess zeta potential. The effect of particle diameter on the radial flow profile is illustrated in Fig. 1.8 for certain typical cases. It is seen that the wall effect increases with the particle

References pp. 37-38

18

Chapter1 '

'

'

i

,

,

,

i

,

,

,

,

,

,

i

[-?

2.5

r

,

,

'

4

..

[ ~ P ~ = L_~c~ = 2 ~ P ~ ' ~ , . j ~

faster at the wall

=mm

o (D > i3. r ~ A L_

i

m

1.5

" 50 bar) to cause an additional pressure driven flow through the column. The resulting mode is called pressure-assisted electrochromatography (PEC) and will be described in section 3 of this chapter. The second approach is an equal pressurization of both inlet and outlet vials with an inert gas at relatively low pressures of 10 to 12 bar as shown in Fig. 2.1. The reasons for formation of bubbles in packed column CEC has not been explained satisfactorily. Bubbles may be formed due to local differences in EOF velocity (e.g. between unpacked and packed sections of the capillary [6,7]), by local differences in field strength (leading to "hot spots"), by release of gas trapped in the pores or electrochemically formed [8]. Whichever mechanism applies, it was suggested by early workers in CEC, to pressurize the inlet and outlet vials, in order to keep the gas dissolved [3,9,10]. Once the bubbles form, the detector base line becomes very noisy and the current unstable. This may lead to break down of the current and the flow stops. Robson et al. illustrated that using pressurization of the solvent vials CEC can be carried out routinely at high fields with high speed and high efficiency [ 11]. Performing reproducible CEC requires stringent control of parameters like temperature, voltage, and pressure. In commercial CE equipment, many of these

Instrumentation

43

parameters are automatically controlled which has led to significant improvements in reproducibility and accuracy of the separations. Currently, most manufacturers of CE instrumentation also provide the option for pressurization of both outlet and inlet vials (see Table 2.1). The Hp3DcE system (now Agilent Technologies Capillary Electrophoresis System, Waldbronn, Germany) has the option to apply a gas pressure of 0.2 to 1.2 MPa to the outlet and/or inlet vial [ 12]. A pictorial view of the system is shown in Fig. 2.2. Isocratic CEC separations are easily performed in this equipment, which accounts in part for the initial success and acceptance of CEC. However, for advanced operation, an extensive modification of the equipment is obligatory and will be described later in this chapter. In a review of the technique, Majors claimed that CEC needs dedicated instrumentation, designed specifically for CEC and not adapted from CE. According to the same author, this was the general consensus in discussions amongst leading researchers in CEC [13, 14]. Shortcomings in current commercial CEC instrumentation include the lack of a reliable instrument allowing (i) gradient elutions and voltages higher than 30 kV, (ii) rapid changes in column temperature, (iii) use of capillaries with varying lengths, and (iv) interfacing to modern sampling formats such as 96 (384) well plates. These shortcomings have led to further development of CEC instrumentation.

2.2.2 Temperature control 2.2.2.1 Thermal effects

The control of temperature is vital in CEC. During the electrochromatographic separation process, electrical energy is released as Joule heat. Part of the thermal energy is dissipated to the environment, while the other part is consumed by the medium resulting in an increase of the temperature in the capillary. The ion mobility depends on the solvent viscosity. Upon the increase in the temperature of the solution, its viscosity decreases causing a further increase in the current, thus releasing more heat (Fig. 2.3) [15]. If the heat dissipation is insufficient, the temperature may raise considerably to a point at which the eluent boils thus forming vapor bubbles. This leads to breakdown of the current. In addition, the increase in temperature results in an unequal temperature across the column diameter, leading to different flow velocities across the capillary cross section, and, consequently to reduction in the column efficiency

[16]. For that reason, as in CE, narrow capillaries with a high

surface-to-volume ratio are used in electrochromatography, so that heat dissipation is more effective. Since the effectiveness of radial heat dissipation decreases with the increase of the capillary diameter, a maximum internal diameter suggested for packed CEC capillaries is 0.2 mm. Obviously, effective temperature control, actually cooling

References pp 83-85

TABLE 2.1 LIST OF COMMERCIAL SYSTEMS USEFUL FOR CEC

Essential specifications

URL

Manufacturer

System

Agilent Technologies, Waldbronn, Germany

Agilent Capillary Forced-air temperature cooled with Peltier element. http://www.chem.agilent.com/cag/ products/hp3dce.html Electrophoresis System Temperature range 10°C below ambient to 60°C (0.1 "C) with a minimum of 4°C. Minimum total capillary length 33.5 cm Minimum effective capillary length 8.5 cm. Vial pressurization to 12 bar. Several modes of operation, CE, CEC, CE-MS, CE + pressure. CEC capillaries 250 x 0.1 mm and 400 x 0.1 rnrn, CIS-, C8- and Phenyl type phases

Beckman Coulter Inc., Fullerton, USA

http://www.beckman.com/ PIACE MDQ Methods Temperature control of capillary column with rapidly beckman/biorsrch/prodinfo Development System recirculating liquid coolant. Selectable temperature control Icapelecl paceseries.asp between and 15°C and 60°C (0.1"C). Solvent delivery provided by both variable pressure (0.1 to 100 p.s.i).

Biomolecular Instruments, Santa Fe, USA

Spectraphoresis Ultra

http://www.biomolecularl .corn/ Peltier controlled 10°C below ambient to 60°C in O.l°C increments. Constant voltage, constant current, voltage gradients, positive or negative polarity, time-programmed polarity switches current gradients, constant power. Pressure at one or both ends of a capillary with the voltage or current can be applied. Minimum capillary length 22.5 cm; maximum length 63.5 cm; shortest detection length ( L d ) 6.3 cm.

5 h

Crystal 300 Series

Constant voltage or constant current, positive or negative pressure, simultaneous pressure and voltage. 10400°C max. (identical with PrinCE System) Pressure: -180 - 250 mbar, 1 mbar increments.

2

$ 9

%

Microtech Scientific &, Inc., Sunnyvale, USA CII

http://www.biomolecularl.corn/

http://www.microlc.com/ Ultra-Plus I1 Capillary Temperature controlled from 5°C above room to 85°C. CE 1 CEC Module Columns 50 pm to 500 pm fused-silica capillary, 1.0 rnm to 2.0 rnm for micro bore. Lenght 1 cm to 100 cm for capillary, 1 cm to 25 cm for micro bore. Flow rate range 0.1-500 pllmin. pressure limit 10,000 p.s.i. Power supply voltage from 0 to 60,000 V.

Unimicro Technologies TriSepTMCEC System Gradient CEC system which includes two PU-980 HPLC Inc. Pleasanton, USA CEC, HPLC and CE pumps, flow rate range 1 pVmin to 10 ml/min, -30 or +30 kv. Electropak CEC columns. ProLab Instuments Evolution 200, GmbH, Kagenstrasse MicroHPLC System 17, CH - 41 53 Reinach, Switzerland

Binary, high-pressure gradient pump. Flow rate range 1-200 pllmin in gradient mode, isocratic 0.1-200 pllmin. Pressure range 0 4 0 0 MPa. Integrated vacuum degasser. Programmable high voltage power supply 30 kV, 200 pA.

http://www.unimicrotech.com/ index.htrn1

Chapter 2

46

"v" t~ ~.,=_

~o t~

"0

~ .Ig e~ o t~

t~

~D r/3 rat3 ~r/3

~E e~

~D

m

el-

O

O

E t_ ,

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O .,..a ~D

-n O t_ t~

~

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!

(D

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(D Cr

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100,000 plates/m. The analysis of bases, however, remains a challenge (see later section). The use of other column chemistries such as C8 and phenyl phases allows the selectivity of the stationary phase to be optimised in a similar fashion to that used in LC [60-62], (see Fig. 3.5).

Referencespp. 106-110

Chapter 3

98 3.4.2 The use of mobile phase additives

As the generation of a significant EOF requires the presence of ionised groups on the surface of the stationary phase, classical silicas with high metal contents have been widely used in CEC. Before "new generation" silica RP column chemistries became available, undesirable secondary interactions of basic analytes in LC were observed. The traditional approach to the analysis of bases by LC using stationary phases containing acidic silanols was to use small competing bases added to the mobile phase, such as triethy!amine or, in CE, triethanolamine has been used. Recent work in this area has demonstrated that strong and weak acids and bases, and neutrals can all be eluted in one CEC run by the use of triethylamine phosphate or triethanolamine phosphate [63] (see Fig. 3.6) or hexylamine at pH 2.5 [64,65]. By the use of this approach acids are chromatographed in their ion-suppressed mode whilst strong and weak bases are positively charged. The uncharged species are separated according to differences in lipophilicity by interactions with the RP stationary phase giving rise to significantly different selectivity to that in CE. The selectivity also differs from LC for the charged bases due to additional differences in electrophoretic mobility. An added benefit is the ability to distinguish bases in mixtures of acids, bases and neutrals as the positively charged species elute before the EOF at pH 2.5. The orthogonal separation mechanism provided by this approach is extremely attractive to the pharmaceutical industry. The inclusion of basic additives in the run buffer leads to a reduction in the EOF. This is due to the reduction in the number of free silanol sites on the silica surface. However, above 50 mM the continued reduction in the EOF is less pronounced [63]. In practice, sufficient EOF is generated, even in the presence of mobile phase additives, to elute neutral species in acceptable times. The upper limit on the additive concentration is most frequently due to excessive baseline noise arising from high background absorbance. The inclusion of mobile phase additives leads to a further level of complexity in method development and prohibits coupling to mass spectrometry. However, this approach is a practical solution until better stationary phases are developed. 3.4.3 Ion exchange - SAX, SCX and Mixed Mode

3.4.3.1 SCX In order to extend the practical working pH range in CEC the use of stationary phases containing charged functional groups has been utilised. A strong cation-exchange phase (SCX, which contains a sulphonic acid group) will be negatively charged from pH 2 to pH 9. Hence, a good EOF is maintained throughout this region

99

Modes o f CEC

140 ~ 120 "~ r~

100

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'

'

I

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2

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4

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Fig. 3.6. Electrochromatogram of benzylamine (I), caffeine (II) and benzoic acid (III). Efficiency values of 4642, 76331, and 6399 plates per column were obtained respectively for I, II, and nI. Electrochromatography was performed at 15~ with an applied voltage of 25 kV on a 25 cm, 100 ~tm i.d., 3 ~tm Hypersil Phenyl packed capillary. Mobile phase: ACN-50 mM triethanolamine phosphate, pH 2.5-H20 (6:2:2 v/v/v). Sample concentration was 100 ~tg ml 1 of each component, 5 kV/5s injection. Detection at 214 nm. From [63]. Reproduced with permission of The Royal Society of Chemistry

[22]. Considerable excitement was generated when Smith and Evans reported efficiencies in excess of 8 x 106 plates m 1 for the analysis of the basic tricyclic antidepressants. These high efficiencies have also been obtained by other workers for a range of structurally diverse basic compounds. Unfortunately these workers, including Smith and Evans, have experienced severe non-reproducibility of the phase, in that severe tailing and fronting have been unexpectedly observed in the middle of successful runs [46,66,67]. Explorations into the focussing mechanism have been reported [68], but it is yet to be fully characterised and understood. However, the separation of weakly basic aromatics on this phase has been demonstrated [21]. An additional disadvantage of the SCX phase is that it possesses little hydrophobic character and hence is not very selective for the separation of neutral analytes. Harnessing the phenomenon of high peak efficiencies in a repeatable fashion could lead to a new surge in the interest for CEC. 3.4.3.2 SAX

In the case of a strong anion-exchange phase (SAX, which typically contains a quaternary amine), EOF reversal is observed, and an ion-exchange mechanism with negatively charged species will also result [69]. This approach has been successfully applied to the analysis of iodide and iodate [70], and has recently been reported for the References pp. 106-110

1O0

Chapter 3

separation of a range of large proteins using a custom synthesised tentacular anion exchanger [71]. The resolving power of CEC with this phase for protein work was demonstrated with separations of chicken egg-white conalbumin variants in a single four-minute run with isocratic eluents. Other workers have reported the use of anion exchanger columns for lanthanide and inorganic ion analysis [29] and aqueous and non-aqueous chiral derivatised amino acid analysis [72,73]. 3.4. 3.3 Mixed mode

There are two types of mixed mode phases 1) physically distinct particles possessing separate lipophilic and ion exchange chemistries [74] and 2) the lipophilic and ion exchange chemistries on the same silica particle [26,75]. The use of mixed mode phases (which contain charged functional groups and lipophilic spacers or separate alkyl ligands) has yielded rapid, efficient methods for the analysis of neutral and ion suppressed acids. Custom synthesised multilayer mixed-mode phases containing ligands with a sulphonic acid sub-layer and a C18 top-layer have also been reported [32,58]. These phases have been successfully used to separate nucleobases and even strongly acidic analytes such as nucleic acids of various sizes from dinucleotides to t-RNAs. Excellent peak area, and migration time precision can be obtained for neutral and acidic compounds for a given mixed-mode column. A major advantage of using mixed mode phases is that they allow the rapid analysis of ion suppressed acidic and neutral analytes at low pH [26]. With the exception of one recent report by Rozing [76], using a new Zorbax SCX/C18, no success has been reported in analysing basic compounds with SCX mixed mode phases. 3.5 CHIRAL STATIONARY PHASES Chiral CEC will be discussed in detail later in the book but is included here to exemplify the application of the high efficiencies obtained with electro-driven techniques which makes them attractive for chiral analysis where selectivity factors are sometimes small. CE has made use of chiral additives in the electrolyte whilst LC tends to utilise chiral stationary phases. Both options have been explored for chiral CEC [27,28,77]. The small amount of packing material necessary for capillaries allows the use of chiral stationary phases that would be prohibitively expensive for standard LC. Cyclodextrins, proteins, antibiotics and molecular imprinting have all been used to form chiral stationary phases [78-80]. After some less than encouraging peak efficiencies obtained using the chiral CEC approach, much improved chiral resolutions have been achieved using CEC compared to LC or CE [81-83].

Modes o f CEC

101

3.6 GEL CEC

Columns packed (filled) with natural and synthetic polymers such as acrylamides, dextrans, glycols, poly(ethylene oxides), poly(ethylene glycols), methacrylates, agaroses and various cellulose derivatives have been widely applied and reviewed for a number of applications in capillary gel electrophoresis [84,85]. The rationale for such a diverse range of polymers is associated with their differences in physicochemical properties which allows separation of a variety of molecules and biopolymers over a wide range of polymer concentrations [86]. Gel CEC is included here simply because many publications describe covalently immobilised polymers or mimetic gels within a fused silica capillary as "electrochromatography" and a distinction between Gel CEC and CGE is therefore difficult to define. Nevertheless, Gel CEC will be covered in greater depth in a later chapter, and it is included here simply because, due to the lack of a clear definition, it could be argued to be the most widely used mode of CEC. 3.7 MONOLITHS The use of hydrothermally formed retaining frits in capillary columns packed with stationary phase particles is an accepted limitation in CEC. The introduction of the frit to hold the packed bed is vital, yet introduces problems such as EOF and flow non-uniformities, compromised frit permeability [87], capillary fragility, increased likelihood of bubble formation [88] and a thermally induced modified frit surface chemistry which can detrimentally alter the chromatography [23]. Practical aspects to be considered include the appreciable effort and skill of the analyst who is required to repeatably manufacture capillaries of a particular phase and redevelop the fritting and packing methodology for each different stationary phase type. The use of continuous bed columns (i.e. monoliths) can address many of these points. Early monolith research focussed on the polymerisation of acrylamides analogous to the approach utilised in CGE [89]. As the area developed, workers tried different chemistries but experienced problems that ranged from shrinkage of the monolith from the capillary wall and / or cracking of the monolith structure - leading to poor chromatography. It is these issues that have generally hampered the development of monolith columns for LC as well as CEC. A monolithic column was recently defined as "A continuous unitary porous structure prepared by in situ polymerisation or consolidation inside the column tubing and, if necessary, the surface is functionalised to convert it into a sorbent with the desired chromatographic binding properties" [90]. Whilst quite general, this definition actually covers a range of methods that can be used to produce continuous bed columns. References pp. 106-110

102

Chapter 3

The many reported methodologies for producing monolithic columns published to date differ in their manufacturing process and the end-product monolith. Porous moulded organic polymers are perhaps the most reported in CEC and involve in-situ polymerisation of monomer solutions in the presence of a porogen using various initiator techniques such as free radical polymerisation [91] or photopolymerisation [92]. By carefully controlling the polymerisation components and conditions for this methodology, monoliths of defined physicochemical properties may be produced which are robust and exhibit excellent chromatographic properties [93]. Particle-fixed continuous beds are another example involving the immobilisation of stationary phase particles of known characteristics using whole column sintering [94]. Finally, sol-gel technology involving the hydrolysis and polycondensation of precursors in a defined solvent in-situ to produce a hydrogel, which may then be converted to a xerogel upon drying before functionalising has been reported [95]. A derivation on this theme is the use of particle-loaded sol-gels where polycondensation of a mixture of polyalkoxysiloxanes and the stationary phase to produce end frits to effectively retain the phase [96] or to completely immobilise the column have been demonstrated [97,98]. Thus, the monolith approach is highly customisable for a particular need or application. It is an important area in column technology development with the number of publications in CEC, ~tLC and even monolith based standard LC increasing dramatically. Details and specific application examples of the use of these various monolith technologies in CEC will be discussed in a later chapter. 3.8 SIZE EXCLUSION CEC In size exclusion chromatography, selectivity for neutral molecules is based on molecular size and shape. The stationary phase consists of either a polymeric gel or a silica gel with controlled pore size. Larger molecules are excluded from the pores whilst smaller molecules, can enter the pores and are hence eluted later. In CEC of charged molecules additional selectivity is introduced based on electromobility. The mobile phase is used to change the molecular shape and/or charge and to optimise secondary interactions. Capillaries packed with unmodified silica gel with pore sizes between 100 and 10 nm have been evaluated for CEC [99]. In addition to the practical application to polymer analysis, the technique is of theoretical interest as a demonstration of the appreciable intra-particle flow in CEC with associated efficiency gains, which is absent in LC. Li and Remcho [ 100] studied packing media pore sizes from 6 to 400 nm and deduced that only the larger pore diameters (>200 nm) supported pore flow, which led to higher peak efficiencies. For SEC a low pore flow is required to obtain selectivity based on exclusion/inclusion in the pore. Hence low ionic strengths must be used to induce double layer overlap in the pores. The optimisation of the

Modes of CEC

103

pore-to-interstitial flow for size-exclusion has been investigated by Stol et al [101 ]. The implications to RP CEC have also been explored [99,102] and the findings indicate that higher efficiencies and linear flow rates are obtained with larger pore sizes (e.g. 400 nm) and higher buffer strengths which favour high intra-particle flow. In further work, Stol et al [103] verified this work and demonstrated substantial intraparticulate flow with much smaller unbonded packing media (30 nm) in addition to the 400 nm material. The high efficiencies observed for the neutral test probes were attributed to better electrokinetic flow homogeneity due to the large pore size and enhancement of the mass transfer kinetics. A rigorous theoretical treatment of flow characteristics in packed beds and determination of an ideal packed capillary structure for electrokinetic flow was offered by Luo and Andrade [104]. 3.9 GRADIENT CEC AND VOLTAGE ASSISTED CAPILLARY LC Preliminary work has been published by a number of workers using research instrumentation [ 105-108]. There are, at time of publication, few commercially available continuous gradient CEC instruments. It is possible to perform simple step gradients using standard CEC instruments by changing the mobile phase during the run [109,110]. This will be a critical area for the development and acceptance of the technique. Preliminary reports using a prototype gradient CEC system capable of performing capillary LC with voltage assisted flow have been very encouraging [25]. It has been found that a large number of compounds of widely different lipophilicity may be eluted and resolved using isocratic CEC that would have required gradients for LC. This presumably is due to the high peak capacity of CEC and hence higher efficiencies compared to LC. The combination of pressure-driven and electro-driven flow offers a great potential for optimisation of separations. For example, CEC systems using columns with low EOF may not elute all species by electromobility alone, whereas the application of pressure will ensure a sufficient flow to elute all species [76,111 ]. The selectivity and structural information afforded by being able to elute analytes with and without applied voltage is also of practical benefit [47]. Once again the availability of commercial instrumentation will be the key to the success of this technique. 3.10 SELECTIVITY COMPARED WITH LC The potential influence of the nature of the driving force on the chromatographic selectivity for neutral molecules has been investigated using CEC, gLC and pressurised flow electrochromatography. The comparison was performed on the same prototype instrument and the same column to eliminate all other possible influences References pp. 106-110

Chapter 3

104

on the selectivity. The relationship between k and the percentage of organic modifier present in the mobile phase was used to compare the consequence of the different mechanisms on the separation of non-ionised molecules. The slopes and intercepts of the plots of log k against percentage organic in the mobile phase, obtained by using the different flow generation mechanisms, were statistically evaluated. As expected, a linear relationship was found between log k and the percentage of organic modifier for a series of weak acids and bases analysed in their ion-suppressed mode and for neutral compounds. Under the conditions investigated, the chromatographic selectivity in electro, pressure driven flow and a combination of thereof was shown to be equivalent for the non-ionised molecules studied [25]. Similar findings were reported by another group in that no significant differences could be demonstrated between CEC, PEC and ~tLC modes on the same capillary for separating a range of 27 neutral analytes [ 112]. In contrast other groups have reported the observation that the chromatographic characteristics of porous reverse phase materials depends on the mode of flow generation [113]. Other authors have additionally reported the different elution profiles between PEC and capillary LC for analytes that include basic amines [47]. On a practical basis, numerous workers such as Ross et al [114] have shown that method transfer between LC and CEC for neutral or ion suppressed analytes is straightforward and simple. The obvious benefits of using CEC compared to LC include increased efficiencies and hence enhanced peak capacity and the orthogonal nature of CEC compared to LC when ionic analytes are present. Retention modelling of neutral [ 115] and basic analytes [12,65] taking into account retention factors and electromobilities of the analytes have shown good agreement between predicted and experiment retention times. These results highlight the predictive nature of CEC and the possibility of performing computer optimisation routines. The possibility of using voltage to "fine tune" pressure separations is an attractive technique which will require more attention when commercially available instrumentation becomes available. 3.11

GUIDELINES FOR THE ANALYSIS OF ACIDIC BASIC AND NEUTRAL COMPOUNDS

3.11.1 Neutral analytes

Traditional LC stationary phase, such as Hypersil or Spherisorb ODS1 materials are ideally suited to the analysis of neutral analytes as the phases possess a high content of acidic silanols, hence at pH 7-9 high EOF generation is achieved which facilitates rapid analysis. In marked contrast, many pharmaceutical compounds pos-

Modes of CEC

105

sess an ionizable functionality and hence different approaches must be used for their analysis as discussed in the sections below.

3.11.2 Acidic analytes Ionised acids tend to migrate towards the anode counter to the EOF therefore they are either not loaded onto the column during electrokinetic injection or are not swept towards the detector and hence are not detected. In order to perform CEC of acidic analytes they must be run in their non ionised form i.e. in acidic mobile phase. As a consequence of the low mobile phase pH there is a significantly reduced EOF hence long retention times are observed. It is highly recommended that the use of mixed mode phases such as the SAX/C18 and SCX/C6 are employed when using low pH mobile phases in order to enhance the EOF [ 116].

3.11.3 Basic analytes The analysis of basic analytes on stationary phases with silica support is still problematic in LC due to the possibility of the basic analyte undergoing mixed mode interactions with the stationary phase i.e. hydrophobic and ionic interactions. Residual isolated silanols are responsible for the deleterious ionic interactions, the result of which is excessive peak tailing. The CEC analysis of basic analytes is thought to be problematic because in order to generate a good EOF, an acidic silica is essential. It is these silanols groups which generate the EOF and are responsible for unwanted peak tailing. A number of approaches to improving the peak shape of basic analytes on silica supports for CEC with stationary phase design have been taken from LC. Many of the problems encountered in the analysis of bases by LC are also manifest in CEC. The use of low pHs or reducing the number of acidic silanols leads to extremely low EOFs which may cause excessive retention of concomitantly chromatographed neutral species in addition to the practical consideration of maintaining a "wetted" capillary. These factors have led to a perception that the analysis of bases by CEC is difficult, if not practically impossible. A number of "new" generation silica support materials have also been evaluated. The reduction in EOF is significant leading to excessive retention time for neutral and acidic species. However, pharmaceutical bases can generally be analysed using such phases, as the intrinsic electromobilities of the molecules are sufficient to ensure elution. Unfortunately, there has been little success to date. The best approach to date for the analysis of basic compounds by CEC has been to incorporate a small basic compound such as triethylamine, triethanolamine [63] or hexylamine [64] into the low pH mobile phase. The small bases act in a competitive manner to restrict the access of the basic analytes to the silanol groups on the surface References pp. 106-110

Chapter 3

106

of the silica. This approach has been shown to work for a number of basic analytes [63-65]. The same approach also allows the simultaneous analysis of acids, bases and neutral, the only drawback being the relative slow EOF. 3.12 CONCLUSIONS The enhanced sample loadability together with the high efficiencies obtainable in CEC have stimulated much interest in this technique. The entire ranges of LC and CE modes are potentially available within CEC. Many of these are only in the initial investigation stages. With the advent of stationary phases specifically designed for CEC and a growing theoretical understanding of the mechanisms involved in CEC, the continued development of the technique is assured. Nevertheless, support from stationary phase manufacturers for custom designed CEC phases and a robust column format are critical for its continued development and acceptance as a mainstream technique. 3.13 ABBREVIATIONS

CEC CGE EOF MES PEC RP SAX SCX SFC TRIS

capillary electrochromatography capillary gel electrophoresis electrosmotic flow 2-(N-morpholino)ethane-sulfonic acid pressure assisted electrochromatography reversed-phase strong anion exchange strong cation exchange supercritical fluid chromatography tris(hydroxymethyl)aminomethane

3.14 REFERENCES

1 2 3 4 5 6

M. Mayer, E. Rapp, C. Marck and G.J.M. Bruin, Electrophoresis, 20 (1999) 43. A. Banholozer and U. Pyell, J. Microcol. Sep., 10 (1998) 321 H. Rebscher and U. Pyell, J. Chromatogr. A, 737 (1996) 171 P.K. Owens and J. Johansson, Anal. Chem., 72 (2000) 740. J. Nawrocki, J. Chromatogr. A, 779 (1997) 29. K.K. Unger, Porous Silica, its properties and use as a support in column liquid chromatography. Elsevier, Amsterdam, 1979. 7 G.B. Cox, J. Chromatogr., 656 (1993) 353. 8 K. Ballschmiter and M. Wossner, Fresenius J. Anal. Chem., 361 (1998) 743. 9 P.K. Owens, O. Gyllenhaal, A. Karlsson and L. Karlsson, Chromatographia, 49 (1999) 327.

Modes of CEC

107

10 B.A. Bidlingmeyer, J.K. Del-Rios and J. Korpi, Anal. Chem., 54 (1982) 442. 11 G.B. Cox and R.W. Stout, J. Chromatogr., 384 (1987) 315. 12 A.P McKeown presented at the Chromatographic Society Spring Meeting, Loughborough, June 2000. 13 A. Maru~ka and U. Pyell, J. Chromatogr. A, 782 (1997) 167. 14 M. Ye, H. Zou, Z. Liu, J. Ni and Y. Zhang, J. Chromatogr. A, 855 (1999) 137. 15 N.M. Djordjevic, F. Fitzpatrick, F. Houdiere and G. Lerch, J. High Resol. Chromatogr., 22 (1999) 599. 16 W. Wei, G.A. Luo and R. Xiang, J. Microcol. Sep., 11 (1999) 263. 17 M.M. Dittmann presented at the 20th International Symposium on Capillary Chromatography, Riva del Garda, Italy, May 1998. 18 P.D.A. Angus, J.F. Stobaugh, C.W. Demarest K.M. Payne K R Sedo L.Y. Kwok and th ' ' " " ' T. Catalano presented, at the 19 International Symposium on Capillary Chromatography and Electrophoesis, Wintergreen, VA, May 1997. 19 W. Wei, G.A. Luo, G.Y. Hua and C. Yan, J. Chromatogr. A, 817 (1998) 65. 20 A.P McKeown, M.R. Euerby, H. Lomax, C.M. Johnson, P. Ross and H. Richie, Chromatographia, submitted, 2000. 21 C.W. Klampfl and P.R. Haddad, J. Chromatogr. A, 884 (2000) 277. 22 N.W. Smith and M.B. Evans, Chromatographia, 41 (1995) 197. 23 R.J. Boughtflower, T. Underwood and J. Maddin, Chromatographia, 41 (1995) 398. 24 K.D. Altria, J. Chromatogr. A, 856 (1999) 443. 25 M.R. Euerby, C. Chevet and C.M Johnson presented at 23 rd International Conference on Chromatography, Granada (1999). 26 M.R. Euerby, C.M. Johnson, S.F. Smyth, N. Gillott, D.A. Barrett and P.N. Shaw, J. Microcol. Sep., 11 (1999) 305. 27 S. Li and D.K. Lloyd, Anal. Chem., 65 (1993) 3684. 28 S. Nilsson, L. Schweitz and M. Petersson, Electrophoresis, 18 (1997) 884. 29 S. Kitagawa, A. Tsuji, H. Watanabe, M. Nakashima and T. Tsuda, J. Microcol. Sep., 9 (1997) 347. 30 M.G. Cikalo, K.D. Bartle and P. Myers, Anal. Chem., 71 (1999) 1820. 31 M.R. Euerby, C.M. Johnson and K.D. Bartle, LC-GC Int., 11 (1998) 39. 32 M.Q. Zhang and Z. E1Rassi, Electrophoresis, 20 (1999) 31. 33 M.M. Dittman and G.P. Rozing, J. Chromatogr. A, 744 (1996) 63. 34 M.R. Euerby, C.M. Johnson, P. Myers, K.D. Bartle and S.C.P. Roulin, Anal. Comm., 33 (1996) 403. 35 M.M. Robson, S. Roulin, S.M. Shariff, M.W. Raynor, K.D. Bartle, A.A. Clifford, P. Myers, M.R. Euerby and C.M. Johnson, Chromatographia, 43 (1996) 313. 36 R.A. Carney, M.M. Robson, K.D. Bartle and P. Myers, J. High Resol. Chromatogr., 22 (1999) 29. 37 R.M. Seifar, W.T. Kok, J.C. Kraak and H. Poppe, Chromatographia, 46 (1997) 131. 38 R.M. Seifar, J.C. Kraak, W.T. Kok and H. Poppe, J. Chromatogr. A, 808 (1998) 71. 39 C. Fujimoto, Trends in Analytical Chemistry, 18 (1999) 291. 40 T. Adam and M. Kramer, Chromatographia, 49(Suppl. I) (1999) $35. 41 T. Adam, S. L~idtke and K.K. Unger, Chromatographia, 49 (1999) $49. 42 T. Tsuda, Anal Chem., 59 (1987) 521. 43 T. Tsuda, Anal Chem., 60 (1988) 1677. 44 J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317.

108

Chapter 3

45 N.W. Smith and M.B. Evans, Chromatographia, 38 (1994) 649. 46 M.M. Dittmann presented at the 18th International Symposium on Capillary Chromatography, Riva del Garda, Italy, May 1996. 47 T. Eimer, K.K. Unger and J. van der Greef, TRAC, 15 (1996) 463. 48 M.M. Dittman and G.P. Rozing, Biomed. Chromatogr., 12 (1998) 136. 49 P.B. Wright, A.S. Lister and J.G. Dorsey, Anal. Chem., 69 (1997) 3251. 50 J.J. Kirkland and R.M. McCormick, Chromatographia, 24 (1987) 58. 51 P. Coufal, H.A. Claesens and C.A. Cramers, J. Liq. Chromatogr., 17 (1993)3623. 52 M.T. Dulay, C. Yan, D.J. Rakestraw and R.N. Zare, J. Chromatogr. A, 725 (1996) 361. 53 P.D.A. Angus, C.W. Demarest, T. Catalano and J.F. Stobaugh, Electrophoresis, 20 (1999) 2349. 54 J. Reilly and M. Saeed, J. Chromatogr. A, 829 (1998) 175. 55 L.H. Zhang, W. Shi, H. Zou, J. Y. Ni and Y.K. Zhang, J. Liq. Chromatogr. Relat. Technol., 22 (1999) 2715. 56 M. Saeed, M. Depala, D.H. Craston and I.G.M. Anderson, Chromatographia, 49 (1999) 391. 57 T. Helboe and S.H. Hansen, J Chromatogr. A, 836 (1999) 315. 58 M.Q. Zhang, C. Yang and Z. E1Rassi, Anal. Chem., 71 (1999) 3277. 59 I.S. Lurie, R.P. Meyers and T.S. Conver, Anal. Chem., 70 (1998) 3255. 60 X. Cahours, P. Morin and M. Dreux, J. Chromatogr. A, 845 (1999) 203. 61 N.W. Smith, CAST, 8 (1999) 10. 62 P.D.A. Angus, E. Victorino, K.M. Payne, C.W. Demarest, T. Catalano and J.F. Stobaugh, Electrophoresis, 19 (1998) 2073. 63 N.C. Gillott, M.R. Euerby, C.M. Johnson, D.A. Barrett and P.N. Shaw, Anal. Commun., 35 (1998) 217. 64 I.S. Lurie, T.S. Conver and V.L. Ford, Anal. Chem., 70 (1998) 4563. 65 M.M. Dittman K. Masuch and G.P. Rozing, J. Chromatogr. A, 887 (2000) 209. 66 M.R. Euerby, D. Gilligan, C.M. Johnson, S.C.P. Roulin, P. Myers and K.D. Bartle, J. Microcol. Sep., 9 (1997) 373. 67 N.W. Smith presented at the 1st International Symposium on Capillary Electrochromatography, San Francisco, CA, August 1997. 68 P.D. Ferguson, N.W. Smith, F. Moffatt, S.A.C. Wren and K.P. Evans, Poster presentation (1999) HPLC 99, Granada, Spain. 69 R. Grtiner, B. Scherer, F. Steiner and H. Engelhardt, presented at the 20th International Symposium on Capillary Chromatography, Riva del Garda, Itlay, May 1998. 70 D.M. Li, H.H. Knobel and V.T. Remcho, J. Chromatogr. B, 695 (1997) 169. 71 J. Zhang, X. Huang, S. Zhang and C. Horwith, Anal. Chem., 72 (2000) 3022. 72 M. L~immerhoferand W.J. Lindner, J. Chromatogr. A, 829 (1998) 115. 73 E. Tobler, M. Lammerhofer and W. Lindner, J. Chromatogr. A, 875 (2000) 341. 74 L. Zhang, Y. Zhang, W. Shi and H. Zou, J. High Res. Chromatogr., 22 (1999) 666. 75 N.W. Smith and M.B. Evans, J. Chromatogr. A, 832 (1999) 41. 76 G.P. Rozing presented at the 13th International Symposium on High Performance Capillary Electrophoresis and Related Microscale Techniques, Saarbrfiken, Germany, February 2000. 77 F. Lelievre, C. Yan, R.N. Zare and P. Gareil, J. Chromatogr. A, 723 (1996) 145. 78 S. Li and D.K. Lloyd, J. Chromatogr. A, 666 (1994) 321. 79 D.K. Lloyd, S. Li and P. Ryan, J. Chromatogr. A, 694 (1995) 285.

Modes of CEC

109

80 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hobo, J. Pharm. Biomed. Anal., 15 (1997) 1351. 81 A. Dermaux, F. Lynen and P. Sandra, J. High Resol. Chromatogr., 21 (1998) 375. 82 C. Wolf, P.L. Spence, W.H. Pirkle, E.M. Derrico, D.M. Cavender and G.P. Rozing, J. Chromatogr. A, 782 (1997) 175. 83 A.S. CarterFinch and N.W. Smith, J. Chromatogr. A, 848 (1999) 375. 84 B.L. Karger, F. Foret and J. Berka, Chapter 13: Capillary electrophoresis with polymer matrices in Methods in Enzymology, Vol 271: High resolution separation and analysis of biological macromolecules Part B Applications., B.L. Karger and W. S. Hancock (Editors) 271 (1996) 293. 85 P.G. Righetti and C. Gelfi, Forensic Science Intl., 92 (1998) 239. 86 A.E. Barron, D.D. Soane and H.W. Blanch, J. Chromatogr., 652 (1993) 3. 87 E.F. Hilder, C.W. Klampf, M. Macka, P.R. Haddad and P. Myers, Analyst, 125 (2000) 1. 88 M.G. Cikalo, K.D. Bartle, M.M. Robson, P. Myers and M.R. Euerby, Analyst, 123 (1998) 87R. 89 C. Fujimoto, J. Kino and H. Sawada, J. Chromatogr. A, 715 (1995) 107. 90 I. Gusev, X. Huang and C. Horvath, J. Chromatogr. A, 855 (1999) 273. 91 J. Liao, N. Chen, C. Ericson and S. Hjerten, Anal. Chem., 68 (1996) 3468. 92 J.R. Chen, M.T. Dulay, R.N. Zare, F. Svec and E. Peters, Anal. Chem., 72 (2000) 1224. 93 N. Ishizuka, H. Minakuchi, K. Nakanishi, N. Soga, H. Nagayama, K. Hosoya and N. Tanaka, Anal. Chem., 72 (2000) 1275. 94 R. Asiaie, X. Huang, D. Faman and C. Horvath, J. Chromatogr. A, 806 (1998) 251. 95 Q.L. Tang and M.L. Lee, J. High Resol. Chromatogr., 23 (2000) 73. 96 M. Schmid, F. Bauml, A.P. Kohne and T. Welsch, J. High Res. Chromatogr., 22 (1999) 438. 97 Q.L. Tang, B.M. Xin and M.L. Lee, J. Chromatogr. A, 837 (1999) 35. 98 C.K. Ratnayake, C.S. Oh and M.P. Henry, J. High Res. Chromatogr., 23 (2000) 81. 99 E. Venema, J.C. Kraak, H. Poppe and R. Tijssen, J. Chromatogr. A, 837 (1999) 3. 100 D.M. Li and V.T. Remcho, J. Microcol. Sep., 9 (1997) 389. 101 R. Stol, W.T. Kok and H. Poppe, presented at the 24th International Symposium on High Performance Liquid Phase Separations and Related Techniques, Seattle, Washington, USA, June 2000. 102 E. Venema, J.C. Kraak, H. Poppe and R. Tijssen presented at the 20th International symposium on capillary chromatography, Riva del Garda, Italy, May 1998. 103 R. Stol, W.T. Kok and H. Poppe, J. Chromatogr. A, 853 (1999) 45. 104 Q.L. Luo and J.D. Andrade, J. Microcol. Sep., 11 (1999) 682. 105 M.R. Taylor and P. Teal, J. Chromatogr. A, 768 (1997) 89. 106 R. Daddoo, C. Yan, D.S. Anex, D.J. Rakestraw and G.A. Hux, LC-GC Int., 10 (1997) 146. 107 C.G. Huber, C Choudhary and C Horvfith, Anal. Chem., 69 (1997) 4429. 108 A.H. Que, V. Kahle and M.V. Novotny, J. Microcol. Sep., 12 (2000) 1. 109 M.R. Euerby, D. Gilligan, C.M. Johnson and K.D. Battle, Analyst, 122 (1997) 1087. 110 J. Ding, J. Szelign, A. Dipple and P. Vouros, J. Chromatogr. A, 781 (1997) 327. 111 T. Eimer, K.K. Unger and T. Tsuda, Fresenius J. Anal. Chem., 352 (1995) 649. 112 Y.K. Zhang, W. Shi, L.H. Zhang and H.F. Zou, J. Chromatogr. A, 802 (1998) 59.

110

Chapter 3

113 J. Jiskra, H. A. Claessens, M. Byelik and C.A. Cramers, J. Chromatogr. A, 862 (1999) 121. 114 G. Ross, M.M. Dittmann, and G.P. Rozing, Publication No. 5965-9031E (1997) Agilent Walbronn, Germany. 115 J.P.C. Visser, H.A. Classens and P. Coufal, J. High Resol. Chromatogr., 18 (1995) 540. 116 M.R. Euerby presented at the 20th Intemational Symposium on Capillary Chromatography, Riva del Garda, Itlay, May 1998.

Chapter 4

Packed Bed Columns Luis A. C O L O N * , Todd D. M A L O N E Y and A d a m M. F E R M I E R t

Department of Chemistry, State University of New York at Buffalo, Natural Sciences Complex, Buffalo, NY 14260-3000, USA ?Present address." The R. W. Johnson Pharmaceutical Research Institute, Science and New Technology~Analytical Development, OMP Bld. B-236, 1000 Route 202, Raritan, NJ 08869, USA

CONTENTS

4.1 4.2

4.3

4.4 4.5 4.6 4.7 4.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Column fabrication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 The column . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Chromatographic material . . . . . . . . . . . . . . . . . . . . . 4.2.2.1 Ion-exchangers and mixed-mode phases . . . . . . . . 4.2.2.2 Submicron particulate materials . . . . . . . . . . . . 4.2.2.3 Highly porous particles . . . . . . . . . . . . . . . . . 4.2.3 Retaining frits . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3.1 Silica-base frits . . . . . . . . . . . . . . . . . . . . . 4.2.3.2 Fritless packed beds . . . . . . . . . . . . . . . . . . . 4.2.4 Fabricating columns . . . . . . . . . . . . . . . . . . . . . . . . Packing methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1 Pressure packing . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.2 Packing with supercritical CO2 . . . . . . . . . . . . . . . . . . 4.3.3 Electrokinetic and pseudo-electrokinetic packing . . . . . . . . . 4.3.4 Packing by centripetal forces . . . . . . . . . . . . . . . . . . . 4.3.5 Packing by gravity . . . . . . . . . . . . . . . . . . . . . . . . . Comparison o f packing procedures . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement ............................. Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

. .

112 112 113

116 129 132 137

139

. . . . . .

140 144 145 150 150 152 152 154 155 156 158 158 159 159

112

Chapter 4

4.1 INTRODUCTION Capillary electrochromatography (CEC) can be performed in open tubes or packed structures. In the open tubular format, the stationary phase is fixed at the inner surface of a capillary column; columns with inner diameter of less than 20 gm are recommended for the best performance [ 1]. Packed structures, on the other hand, consist of a capillary tube filled with chromatographic media. These packed structures can be classified into three different groups: 1) columns packed with particles [2-30], 2) columns containing separation material that has been polymerized in situ, creating a "rod-like" monolithic structure also known as continuous beds [31-39], and 3) columns with entrapped particulate material, which are a combination of the first two groups [40-46]. Columns of the first group are the ones used the most in CEC and will be discussed in this chapter; the other two groups are discussed in Chapters 5, 6 and 7. The chromatographic packing material most commonly used in CEC is HPLC reversed-phase type on spherical particles (1.5-10 gm diameters); although new alternatives are being explored to fabricate materials with more applicability to CEC. As the CEC column technology develops, however, the preference of using columns packed with particles may change, particularly with the emerging approach of monolithic columns. Several protocols can be used to fabricate packed bed structures for use in CEC. In this chapter, we will discuss the packing techniques and column fabrication protocols that have been used for packing particulate material. We concentrate, therefore, on the different approaches used to deliver chromatographic particles into the capillary column. We present an overview of the different packing protocols available to the practitioner, as well as of the CEC column fabrication method, as performed in our laboratory. Our own experiences, practices, and views regarding packing procedures are also provided, when appropriate. 4.2 Column fabrication

Despite the several detailed procedures reported for the fabrication of packed columns for CEC [ 14,17,20,27,30,47-50], column fabrication may still be regarded as an art. A reliable and reproducible performance of a column depends on the column fabrication. Poorly packed columns can lead to low efficiency, poor resolution, and asymmetric peak shapes. The capillary tubes typically used to fabricate CEC columns are fused silica tubes with inner diameters of 100 gm or less, with 50 and 75 lam I.D. being the most popular. The small inner diameter allows for heat dissipation, which is generated by the applied electric field. Packing such columns is an elaborated process and a skill that requires experience.

Packed Bed Columns

113

4.2.1 The column

A packed column in CEC consists of two segments- a packed and an unpacked (or open) section, in most cases. A typical CEC column with a packed and an open segment is illustrated in Fig. 4.1. Most frequently, capillary tubes with less than 100 ~tm I.D. are packed with reverse phase HPLC materials of 1.5-10 ~tm diameter. The chromatographic material is kept in place by means of retaining frits (vide infra). The electroosmotic flow (EOF) velocity in each segment of the CEC column is different [51 ]. The overall EOF velocity depends on the fraction of the packed segment [51,52]. The resulting net EOF is thus a combination from both, the packed and the open segments. To facilitate detection through the column by spectroscopic means, the polyimide coating on the open segment, close to the outlet retaining frit, is removed; this provides the optical window for detection. This can be achieved by any of the methods already in use for capillary electrophoresis (CE) [53]; burning off the polyimide coating is the most common approach. Because of light scattering by particles, optical detection through the packed bed has been reported to decrease detectability [18,24,27]. The length of the column can alternatively be packed completely with the desired separation material (no open segment); however, if detection through the packed bed is not performed, connecting tubing to a detection system is required. Fig. 4.2 shows an example of a 75 ~tm I.D. fused silica column completely packed connected to a piece of a 50 ~tm I.D. capillary used for detection. The butt connector is made by inserting the two capillaries into a piece of PTFE shrinking tubing, which upon application of heat secures the two capillaries in place [54]. It has been reported that butt connection of capillaries has an insignificant contribution to band spreading [55]; however, care must be exercised since connecting of two different pieces of tubing

Packed Section

Open Section

..J

I ~176

""

"-'-'

.. . . . .

i

I Detection Window

Retaining Frits Fig. 4.1. Schematic of a typical packed-capillary column for CEC, illustrating the open and packed segments. References pp. 159-164

114

Chapter 4

Fig. 4.2. Photograph of a butt connection between a 75 ~tm I.D. packed fused silica capillary and a piece of a 50 ~m I.D. capillary tube. Reprinted from ref. [54] with permission. Copyright Wiley-VCH 1999.

2.0

i y

.

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-

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/ 1.4

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i 0.20

,

I

0.30

,

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0.40

,

| 0.50

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Fractional length of the capillary segment packed with bare silica

Fig. 4.3. EOF mobility as a function of the fractional length of the bare silica packed segment for a 100 p,m I.D. capillary containing a 20 cm ODS segment. Reprinted from ref. [56] with permission. Copyright Wiley-VCH 1999.

always has the potential of introducing band broadening to the system. Nonetheless, this option is often used to connect detection schemes that offer higher detectabilities, such as Z-cell for UV detection, mass spectrometry (MS) and NMR detection schemes (see Chapters 2 and 8). In such instances, the gain in detectability and/or structural information is far more important than the loss in efficiency. Totally packed capillary columns, having one segment packed with the stationary phase and a second segment with bare silica, have been fabricated to control the EOF [56]. In this case, the segment that is open in a typical CEC column is packed with bare silica to accelerate and provide a steadier EOF. Such a configuration has allowed an increased EOF that translates into shorter analysis times. Fig. 4.3 and 4 show the EOF mobility as a function of the fractional length of a column packed with bare silica and the effect on analysis time, respectively. As the porosity of the bare silica particles is increased, the EOF is also increased [56]. Columns have also been packed

Packed Bed Columns

115

'i 5

a, u = 0.8 mm/s Nay = I01,000 plates/m E tt)

b.

~g

u = 1.0 mm/s Nay = 108,000 platefdm

2

"~'5

C, u = 1.1

mm/s Nay = 130,000 plates/m

r

2'0

Min

Fig. 4.4. Electropherograms illustrating the effect of the length of the bare-silica segment on the separation of probe compounds, a) 0 cm, b) 6 cm, and c) 28 cm. Solutes: 1, benzene; 2, toluene; 3, ethylbenzene; 4, propylbenzene; 5, butylbenzene and 6, pentylbenzene. Reprinted from ref. [56] with permission. Copyright Wiley-VCH 1999.

with a blend of bare silica and reverse-phase silica supports [18,57]. This can provide enhanced EOF due to the amount of silanols groups introduced by the bare silica, decreasing analysis time. This approach also reduces retention because of the decrease amount of stationary phase, as the bare silica replaces the bonded one; hence, retention depends on the blend ratio. Initial work on CEC was performed on drawn-packed capillaries [ 11 ], a procedure originally introduced by Tsuda et al. [58]. In this approach, large bore columns (thick walled Pyrex tubing) were packed with underivatized packing material; then the columns were pulled at high temperatures to a desired diameter using a glass drawing machine. The stationary phase was attached to the underivatized support packing material after the columns were drawn. This column preparation procedure is not currently used because of the low success rate in fabricating the columns.


116

Chapter 4

4.2.2 Chromatographic material Because of the column's dimensions in CEC, it is important to consider a narrow size distribution of the particles. The effect of particle size distribution on separation efficiency in CEC is expected to be similar of that in HPLC. Although of the same nominal particle size, different packing materials can yield different efficiencies in CEC. It has been pointed out that the different efficiencies reported for the separation of polycyclic aromatic hydrocarbons (PAH), for example, using different packing materials of the same sizes, can be attributed in part to the size distribution of each material [59]. The structure of the packed bed can be influenced by the size distribution. A homogeneous packing size leads to well-packed beds, approaching a closed packed structure. This can be seen in Fig. 4.5, where panels A and B show SEM of the packed bed for columns that were packed with silica particles of about 3 and 0.5 ~tm in diameter, respectively. It is apparent that the particle size in panel A is not as homogeneous as that of the particles in panel B. Notice how the particles with the tighter size distribution form a better-packed bed. One of the most important properties of a column packing material for CEC is the ability to support EOF. This is not only necessary for the separation of neutral compounds but also to separate charged species as the EOF is responsible for the bulk transport of the mobile phase and analytes [7,16]. In the absence of EOF, only species with the appropriate charge will reach the detector. Therefore, packing materials with very favorable characteristics for EOF generation are desired in CEC. Many silicabase HPLC packings from 1.5 to 40 ~tm in diameter have been utilized to pack columns for CEC; those with the C18 reverse- phase being used the most. Other materials include C8 [60-62], phenyl [61,62], and C30 [63,64]. The surface silanol groups of the silica impart a negative charge to the packing material, leading to the generation of the EOF upon application of the electric field. The high surface area of the silica-based packed beds provides for the EOF to be generated mostly at the particle surface, with negligible contributions from the fused silica capillary walls [7,16,65]. However, it is apparent that not all C 18 reversed phase HPLC materials are suitable for CEC. For example, Table 4.1 summarizes electrophoretic mobilities observed on typical reverse-phase HPLC chromatographic materials, and Fig. 4.6 illustrates the CEC separation properties of several C 18 packing materials under identical separation conditions. Faster analysis times are achieved with those materials capable of generating a strong EOF. CEC Hypersil and ODS-1 type are popular among the reverse-phase materials since they seem to support the fastest EOF. These materials are HPLC supports that have not been end-capped, and therefore, a relatively large amount of silanol groups are left on the surface, which can generate EOF. The EOF decreases as the alkyl substitution at the packing surface is increased be-

Fig. 4.5. SEM of the packed bed for columns packed with silica particles of about (A) 3 and (A) 0.5 prn diameter, respectively.

118

Chapter 4

TABLE 4.1 ELECTROOSMOTIC MOBILITIES OF VARIOUS CHROMATOGRAPHIC MATERIALS UTILIZED FOR CEC

Stationary phase material

Electroosmotic mobility (x 10-4 cm2/Vs)

BDS-ODS Hypersil a

0.99

CEC Hypersil C 18a

2.26

Hypersil ODS b

0.14

LiChrospher RP-18 b

1.45

Nucleosil 5 C18 b

1.56

ODS Hypersil a

1.47

Partisil 50DS3 b Prontosil polymeric C30 c Purospher RP- 18b

205) in order to minimize parabolic flow [110]. Under such conditions, 80% of the volume transport due to electroosmosis should be retained [ 110]. Overlapping of the electrical double layer becomes significant in channels with diameters less than 206 and the plug-like profile is lost, decreasing EOF considerably. The double layer thickness has typical values of less than 10 nm, depending on the electrolyte concentration. The thickness of the double layer as a function of acetonitrile concentration at different ionic strengths, using univalent ions, is shown in Fig. 4.15. To at least maintain 80% of the flow transport for the double layer thickness in the typical range of 1-10 nm, the minimum flow channel dimensions must be 20-200 nm. In CEC, the EOF depends on the column packing structure and pore size of the packing material [51,111-116]. In a packed bed, there are many interconnected channels between particles, which leads to a porous packed structure. The porosity of the packed bed dictates the permeability through the column. The average channel size between particles in a CEC column can be estimated if the packed bed is assumed a


Chapter 4

134 --- 40

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Percent of Acetonitrile Fig. 4.15. Thickness of the double layer as a function of acetonitrile concentration in a water-acetonitrile mixture, as per Equation 4.1. Constants for the mixed solvents were obtained from ref. [49,129].

collection of capillary tubes with an average diameter corresponding to the channels between particles. Equation 4.3 shows a relationship between the mean channel diameter and particle size (alp), accounting for the particle structure through the interparticle porosity [ 113]:

dc = 0.42 alp g 1-e,

(4.3)

where ~ is the interparticle porosity. A fairly well packed column is considered to have a random packing structure with an interparticle porosity of 0.4 [117]; therefore, the channel diameter is given by:

dc- 0.28 dp

(4.4)

A similar channel diameter (i.e., 0.25 dp) was originally suggested by Knox who

Packed Bed Columns

3.530 ~o

2.5

135

9

~

................ --m---- 3 pm

i

iiiiiii

--T--0.2

2.0 O

o

~.5

LL

1.0

0 LU

0.5 00

: ..................... !................. ~;~:: ..........

t

.......... i"
20 8. EOF linear velocities above 2 mm/s have been achieved in CEC using packed beds with submicron particles and organic/aqueous mobile phases [66,120]. So far, the separation efficiencies reported with the submicron packed beds have not offered a significant improvement over those obtained with particle diameters in the 1 ~m range [66,119-121]. Fig. 4.17 depicts the separation of a test mixture obtained in a packed bed with particles of about 0.5 ~tm in diameter. As reported by Luedtke, et al. [ 121 ], plate heights of about three times the particle diameter (H = 3dp) are achieved. This has been attributed to band dispersion due to temperature effects and instrumental limitations, such as the maximum electric field that can be applied with existing units and detection systems [121]. Plots of plate height versus linear


Chapter 4

136

1) Thiourea

2

2) Naphthalene 3) Ethylnaphthalene 4) Amylbenzene

I

I

I

0.4

I

0.8

I

I

1.2

I

1.6

Time/minutes Fig. 4.17. Separation of a test mixture in a packed bed of 0.5 ~tm (C8) particles. Column: packed bed of 15 cm in a capillary of 35 cm total length; mobile phase: 80:20 acetonitrile-50 mM Tris buffer at pH 8; separation voltage of 30 kV; injection, 3 s at 300 V; UV detection (220 nm).

velocity indicate that a minimum in plate height has not been achieved and higher electric fields are needed to achieve higher velocities (see Fig. 4.18). Nevertheless, the small particles do provide for rapid separations with current systems. Further studies are still required to obtain a complete understanding of the effect of the submicron material in CEC.

137

Packed Bed Columns

3.5 (# c

.~

~, |

3.1

~ X

~ ~

~1'~" C

E v

=L

F- 2.7 "1-

T

(9

0.2

0.6

w -1-

1.0 1.4 Ttme/mln

1.8

LU 2.3 F< J &.

1.9

1.5 ,

0.1

I

,

0.6

I

1.1

~

I

1.6

,

I

2.1

,

I

~

2.6

I

3.1

LINEAR VELOCITY (mm/s) Fig. 4.18. Plate height versus linear velocity for 9-(1-pyrene)nonanol, last eluting peak in electropherogram in the insert, obtained in a column packed with particles of about 0.5 ~m diameter. Column: 12 cm packed, 35 cm total length; mobile phase 80:20 acetonitrile-50 mM Tris pH 8; voltage 27 kV.

4.2.2.3 Highly porous materials

The typical porous silica-base materials commercially available have pore sizes close to 10 nm. Based on our discussion on the channel diameter requirements to support EOF, it is unlikely that EOF through such pores can be generated under normal CEC conditions. However, it is evident that intraparticle EOF exists in materials with relatively

large pores,

exhibiting perfusion

through

the particles

[45,56,116,122,123]. Flow transport through a highly porous particle is schematically represented in Fig. 4.19. With appropriate channel diameters, EOF can be generated within the particle, transporting the solute through. In pressure driven systems, there is no flow through the particle pores and solutes can only have access to the pores by diffusion. Perfusive transport through wide-bore silica particles with nominal pore


138

Chapter 4

Deft,~;~',~.~%~, ---;--

Dapp~, ~:. ~?~"'"

9

(a) Pressure-drivenFlow (HPLC) Diffusion only

~o

[:::)m

..~,:~ .......

'" "'~ "

'

(b) Voltage-drlven Flow (CEC) Dlffuslon + Convection

Fig. 4.19. Schematic representation of intraparticle flow. In pressure driven flow there is no flow through the particle (A); in electrically driven flow there is intraparticle transport. In (A) transport of solute into the pores is accomplished solely by diffusion, whereas in (B) the EOF enhances transport through the pores. Reprinted from ref. [125] with permission. Copyright Elsevier 1999.

sizes larger than 200 nm was initially reported by Remcho and coworker [ 123]. Later Stol et al. showed intraparticle EOF in particles with pore sizes between 50-400 nm [122,124]. The EOF transport through wide-pore silica has been reported even in particles entrapped in capillary columns via sol-gel processing [45]. Double layer interactions within the pores are minimized by controlling the concentration of the electrolyte in the mobile phase. EOF through the pores is observed at high concentration of electrolyte, while at low concentration of electrolyte, double layer interaction occurs and transport through the pores can be stopped [122,123]. The effect of electrolyte concentration on separation efficiency for columns packed with 7 ~m (C18) particles containing pores with a diameter of 400 nm (nominal value given by manufacturer) is illustrated in Fig. 4.20 [122]. The flow through the pores provides for an enhanced mass transfer, resulting in improved separation efficiencies. Separation efficiencies of 430,000 theoretical plates/rn have been reported for the columns packed with the 7 ~m (C18) particles having 400 nm pore diameter [122]. The efficiencies and linear velocities observed with the highly porous packing materials are comparable to those obtained with small particle sizes. Therefore, the use of the highly porous large packing materials have been proposed as an alternative to the very small particles to perform fast separations with high efficiency in CEC [122,123,125]. The wide-pore large particles also offer the advantage of easier packing than submicron material, since aggregation is minimal. However, highly porous particles have the disadvantage of being fragile; hence, care must be exercised during packing to avoid damage of the particles. The enhanced particle porosity will also affect sample capacity adversely and the increase in ionic strength to maintain a thin double layer can lead to heating of the column.

Packed Bed Columns

13 9

10

7.5

H (pm) 5

2.5

0

1

2

3

4

u (mm/s) Fig. 4.20. Effect of the electrolyte concentration in the mobile phase on separation efficiency in a packed bed containing 7 pm highly porous C18 particles (Nucleosil 4000-7). Reprinted from ref. [122] with permission. Copyright Elsevier 1999.

4.2.3 Retaining frits The flits retaining the chromatographic packing material inside the capillary column in CEC seem to be the "Achilles heel" of the packed column fabrication process. They are the major problem in column manufacturing and perhaps the most critical parameter influencing column performance in general [5,126,127]. Most typically, the flits are fabricated by sintering silica-base packing material by means of heating. Using this approach, the CEC column becomes fragile at the flit since during its fabrication the protective polyimide coating is removed. There also seems to be lack of reproducibility and reliability in the manufacture of the flits, particularly between laboratories. The characteristics of the packing material at the flit position change as heat is applied to produce the flits. This creates non-homogeneous packing at the flit, having different electrical resistivities when compared to the open and pack segments of the columns. The different electrical properties of the flit can contribute to non-uniformities in EOF, and lead to bubble formation at the boundary between the flit and the unpacked segment of the capillary [ 10,51,127]. Constructing flits with resistivities similar to either the packed or the open segment can minimize discontinuities in the column structure, hence decreasing non-uniformities in EOF. Bubble formation, which has also been attributed to heat generation as the electric field is applied, can be reduced by several means. The most common approach to avoid bubble formation is to pressurize the mobile phase at both inlet and outlet column reservoirs References pp. 159-164

140

Chapter 4

[ 10,14,17,27,128]. Other common practices include the use of well-degassed solvents, low concentrations of electrolytes, a relatively large amount of the organic component in themobile phase, working at reduced temperatures (e.g., 15~ when possible, and the use of low conductivity electrolytes (i.e., zwitterionic buffers). The addition of sodium dodecyl sulfate (SDS) into the mobile phase at low concentrations has also been used to minimize bubble formation [67]. The effects that frits and packing materials have on EOF seem inconsistent. For example, placement of two frits in a capillary column (without chromatographic packing material) has shown to be flow restrictive points, reducing EOF by 35% compare to a capillary without frits [56]. On the other hand, an increase in EOF has been reported for a column packed with ODS material when compared to an equivalent open tube [90,129]. The discrepancies can be attributed to differences in the materials and procedures used to fabricate the frits, since these appear to be the major differences reported [ 130]. The bed-retaining frits must posses high permeability to solvent flow, yet the flits must be mechanically strong to retain the packing material and resist the pressures used to pack and/or rinse the column. The heating conditions and method used to prepare the flits affects such characteristics. Pressure resistance studies have shown that, within certain constrains, correlation between pressure resistance of a frit and its influence on the EOF is likely to be insignificant [ 130]. 4.2.3.1 Silica-base frits

The fabrication of frits has been studied in detail by several researchers [27,55,56,129-131 ]. Behnke, et al. [27] studied the performance of columns fabricated using three different frit fabrication procedures. In one of the procedures, the frits were constructed by sintering (using heat) a plug of silica gel wetted with potassium silicate. The frits were mechanically stable; however, under CEC conditions the columns with these frits showed baseline and electrical current stability problems. Another procedure used the method by Cortes, et al. where the flits were formed by polymerization of a potassium silicate solution containing formamide [132]. The columns fabricated using these frits suffered from similar problems. The third method involved the sintering of a silica gel plug wetted with water. Packed beds retained by these frits showed a stable baseline and current. However, they lacked mechanical stability with relatively large column diameters (150 ~m I.D.); decreasing the column diameter to 50 ~tm I.D., increased the stability of the frit. In a different study, Chen and co-workers optimized the silicate polymerization method [ 131 ]. In their approach, the outlet frit is prepared by first filling the column with a sodium silicate solution. Then, the portion of the column at which the frit is desired is brought in contact to a heating element for a few seconds and the frit is

Packed Bed Columns

141

formed. The polyimide coating is not removed under the heating conditions, reducing column fragility at the flits. The excess of the silicate solution is removed by pressure. The flit is then silanized with a solution of 0.02 mol/1 trimethylchlorosilane in DMF, using imidazole as an acid acceptor. The column is packed and the inlet frit is constructed by a quick dip of the column entrance in another silicate solution and heating. Optimal sodium silicate solutions were found to be 10.8% and 5.4% (w/v) for outlet and inlet flits, respectively. The method uses short heating times: 5-6 seconds for the outlet flit and about 1 second for the inlet one, producing relatively short flits (-~75 ~tm). Columns using these flits have been run without pressurization and showed to be mechanically stable without electrical current stability or bubble formation problems over a wide range of acetonitrile-water mixtures, even under relatively high currents conditions (-27 ~tA) [ 131 ]. The most commonly used approach to fabricate silica-base flits, however, is to sinter the actual chromatographic material in place. Care must be taken, however, in order to minimize degradation of the alkylated silica that will not be part of the flit [24]. Heating time used for sintering depends on column I.D., particle size, and type of stationary phase material to be fritted [133]. During heating, a substantial number of silanol groups can be created, changing the surface charge of the material at the flit [ 130]. Excessive heating also damages the outer side of the capillary column, which can create possible adsorption sites. Fig. 4.21 illustrates the effect of using excessive heating to fabricate the flit. Panel A in Fig. 4.21 shows a magnification of the outer side of the column and panel B shows the entire end of the column. Fig. 4.22 shows an adequate sintered frit. It has been shown that decreasing the created silanols by resilanizing the flits can reduce the likelihood of bubble formation, creating a uniform structure that resembles the packed bed, and leads to an improved separation [55]. Fig. 4.23 depicts electroctrochromatograms for the separation of a test mixture on 3 ~tm Spherisorb ODS 1 packed beds (a) before and (b) after resilinazing the outlet frit with ODS. Silanization deactivates undesirable adsorption sites at the flit with which the analytes can interact [ 134]. The packing functionality can also affect the final properties of the flits. This has been shown on flits formed in open tubes using different silica with different functionalities: bare silica, strong cationic exchanger (SCX), strong anionic exchanger (SAX), and Hypersil ODS silica, without the contribution of any other packing material present [130]. The EOF velocity obtained with each flit is different, depending on residual groups of the different packing. Frits formed with bare silica and SCX showed EOF velocities higher than an open tube and flits of ODS material. The amount of residual groups on the surface of the silica support depends on the heating time applied during the formation of the flit, and can contribute considerably to the overall flow. In a column containing a small flit formed with SAX material a reversal


Fig. 4.21. Effect of excessive heating on the capillary column when fabricating a retaining frit, (A) outer surface of the capillary and (B) entire end of the capillary.

g

42 Y

Q

143

Packed Bed Columns

A

13

Fig. 4.22. Adequate frit without deformities on the capillary, particles are 1 pm in diameter.

of the EOF has been observed, when compared to that in an open tube, under otherwise similar experimental conditions. The temperature of the filament used to fabricate such a frit was 430~


and heating time between 12 and 15 seconds. If the

Chapter 4

144

Before rccoating the fdt

(a)

e

t ,oo-{

b a,

400

I] II

d

em

8l~

1000

1200

Ttme / M.mutes

a Dimethyl phthalate b niU'obmzcne

1400

(b) After recoating the frit a

l l

I

x

5/)0

11 I [

I I

lO~X)

e

{~,_ D~.e~hylph~d~ a

II b

I

15oo

~obe~.e

It c. anisole

~1)o

2~/)o

Tune / 1,~u~es

Fig. 4.23. Electropherograms of a test mixture obtained in 3~m Waters Spherisorb ODS-1 packed bed (a) before and (b) after recoating of the outlet frit with ODS; mobile phase, 70:30 acetonitrile-water. Reprinted from ref. [55] with permission. Copyright Wiley-VCH 1999.

heating time is beyond 15 seconds, the EOF towards the positively charged electrode is reduced considerably, indicating a reduction of the residual positive charge responsible for the EOF reversal [130]. It is important to realize how critical is the heating during frit fabrication, as illustrated with Figs. 4.21; the process must be optimized, requiring experienced individuals. An alternative to sintering frits, which deserves mention here, is to form frits via UV photopolymerization of a glycidyl methacrylate and trimethylolpropane trimethacrylate solution (UV radiation, 365 nm for 1 hour) [135]. The photopolymerization process is similar to that used in the fabrication of monolithic columns (Chapters 5 and 6). Frits fabricated with this method have shown to be reproducible; since there is no sintering of packing material, weakening of the capillary column by removal of the polyimide coating and/or alteration of the stationary phase at the frit are avoided.

4.2.3.2 Fritless packed beds As an alternative to the formation of frits, the packing material can be retained by means of tapers fabricated on the fused silica column. In such a case, the packed bed is completely fritless. Two types of tapered capillary columns have been prepared for CEC: internally [136,137] and externally [133,138] tapered, which have been shown

Packed Bed Columns

145

A

B Fig. 4.24. Schematic representation of (A) external and (B) intemal tapers at the outlet of the CEC column. Adapted from ref. [ 136] with permission. Copyright Elsevier 1998.

to be useful in coupling CEC with mass spectrometry and NMR (Chapters 2 and 8). A schematic representation of the tapers is shown in Fig. 4.24. The internally tapered columns are fabricated by sealing the end of the capillary by means of a high temperature flame; the sealed end is carefully grinded to produce a small opening. A laser-based micropipette puller is utilized to make the externally tapered columns. In an attempt to have a fritless column, a capillary having an external taper was packed and the tapered end served as the entrance to the column; no outlet flit was constructed [133]. The problem with these columns is that they can only be used if the electrophoretic mobility of the packing material is larger than the EOF generated, so it can remain in place. Even in such cases, it can be problematic to keep the packing material inside the capillary. The external tapers are weak points on the column, prone to breakage; hence, externally tapered columns are inherently fragile. The internally tapered columns, on the other hand, are not fragile since only the inner diameter is reduced in dimension not the outer diameter and are easily connected to other tubing via shrink tubing. A manufacturing procedure for an internally tapered column is shown in Figs. 4.25 and 26. In Fig. 4.25, the internal taper is prepared at the outlet end of the packed bed, using a sintered frit at the entrance of the column. Fig. 4.26 describes the procedure for a fritless column. The internally tapered capillary columns can also facilitate the procedure of column packing by obviating the need for having a temporary retaining frit (vide infra). This approach holds great promise since it eliminates the problems associated with frits. Because of the connecting tubing, there is potential for band broadening, although it can be minimized.

4.2.4 Fabricating columns The most commonly used procedure to fabricate a packed capillary column for CEC is depicted in Figs. 4.27 A and B. Although several laboratories may have slight variations, the general procedure is as follows. A piece of fused silica capillary of a


Chapter 4

146

Preparing the tapers A e.g. 60 cm length of fused silica capillary is sealed at the middle with a microflame torch.

fused silica capillary

The seal is cut to yield two tapered capillaries. Non-tapered end to be coupled are ground plane and smooth with P4000 (wet). Tapers are ground to form the desired orifice (i.d. 10-50 lam).

Coupling the segments dead volume free

internal tarter

The ground ends are carefully aligned and pushed together. The dual PTFE/FEP-connector is shrunk onto the junction. Slurry preparation 10 - 20 mg beads are ultrasonicated for 20 minutes in 70 - 150 lal acetone (or methanol). Packing the capillary The slurry is flushed in.

The stationary phase beads are allowed to settle under ultrasonication. Pressure drop to zero over a period of 20 min. The capillary is flushed with water.

e)

dual PTFE/FEP shrink-tube-connector

AP = 500 bar b.~ slurry

30 min. sonication AP = 500 bar acetone (resp.methanol)

30 min. sonication AP = 500 bar H20 r

Frit sintering (T=500~

AP = 500 bar

h~ p~

H20 f)

Burning the detection window

g)

Conditioning

inlet flit Column is flushed for 20 minutes with mobile phase (AP = 150 bar) followed by Electrokinetic conditioning:45 rain. at 10 kV with a 25 min. voltage - ramp 45 min. at 15 kV with a 5 rain. voltage - ramp g)

Storage Column is flushed for 30 rain with iso-propanol (AP = 150 bar) Capillary is stored with both ends immersed in iso-propanol filled vials

h)

~

[

UV - window

/

Internal taner

.............

25cm

i

0.5-2cm

i

lOcm

i

Fig. 4.25. Procedure to fabricate a single-flit CEC capillary column. Reprinted from ref. [137] with permission. Copyright Elsevier 2000.

desired length is selected, usually about 10 cm longer than the actual column length wanted. Usually, the capillary tube is rinsed before use; it is our practice to rinse the tube with a sodium hydroxide solution (-0.1 mol/1) and then water. A provisional frit is sintered at one end of the capillary tube. Typically, this is accomplished by tapping onto a pile of wet silica material, allowing a small section of the column end to pack;

Packed Bed Columns

147

fused silica capillary

a) Preparing the tapers

b)

c)

Slurry Preparation

internal taper

Packing the capillary

AP = 500 bar

The slurry is flushed in.

~,~

slurry

The stationary phase beads are allowed to settle.

60 min sonication AP = 500 bar

Pressure drop to zero over a period of 20 rain.

acetone (resp. methanol)

Dry out overnight. d)

dual PTFE/FEP shrink-tube-connector

Coupling the segments dead volume free The ground ends are carefully aligned and pushed together. The dual PTFE/FEP-connector is shrunk onto the junction.

d)

Burning the detection window

e)

Conditioning

Storage g)

T w o - peace Fritless Column internal taper

i

i)

I

25cm

2cm

UV - window

[

10cm

I

T h r e e - peace Fritless Column int~rnnl tarter

I

25cm

internal tarter

I

1-2cm

U V - window

I ~

I

,0om

I

Fig. 4.26. Procedure to fabricate a fritless CEC capillary column. Reprinted from ref. [137] with permission. Copyright Elsevier 2000.

then heat is applied to sinter the material. The amount of heat required to form the flits depends on the column diameter and particle diameter [133], as well as the heating element used. Different heating elements have been used and they vary from


Chapter 4

148

A

B

,

C

D

>

===C,

.

.

.

.

.

.

_1 .

.

.

.

.

/" wet silica

heating element

frit

flush

Fig. 4.27A. Representation of the steps involved in the column fabrication processes: (A) introducing with wet silica material into the end of the capillary, (B) the silica material is ready to fabricate the temporary frit, (C) fabrication of the temporary frit with a heating element, and (D) excess of silica material is flushed out after temporary frit is made.

optical splicers [28,123] thermal wire strippers [28], microtorch [18], burners [56], to heating elements incorporated into more sophisticated assemblies [ 14]. One relatively inexpensive setup used in our laboratory, makes use of a soldering gun fitted with a Nichrome ribbon (1 mm thick) as the heating element. The ribbon is punctured making a small hole (-0.5 mm) through which the column can be inserted; this facilitates heating of a small spot at any desired point of the capillary. Once the temporary frit is in place, the column is flushed to remove the excess material used to fabricate the frits. The mechanical stability of the frit can be tested by applying pressure using a HPLC pump. The frit must resist the packing conditions; yet, it should be permeable enough to allow solvent flow. The temporary flit is eventually removed. An alternative to the temporary frit is to connect the end of the capillary column to a union containing a metallic frit; this will retain the packing material inside the column during the packing process. The capillary is then packed to a desired length (vide infra). Once packed, the capillary column is rinsed with water,

149

Packed Bed Columns

-

2nd

1st

~ o...

m

o >

12

o q., r

c

. m

o

,

,

i

|

,

,

|

,

lo'oo . . . .

Pore size, nm

ls'oo

Fig. 6.31. Effect of mode pore diameter on flow velocity of the mobile phase through monolithic capillary columns. (Reprinted with permission from [55]. Copyright 1999 Wiley-VCH). Conditions: stationary phase poly(butyl methacrylate-co-ethylene dimethacrylate) with 0.3 wt. % 2-acrylamido-2-methyl-l-propanesulfonic acid; Column 100 ~tm i.d. • 28 cm; mobile phase, 75:25 vol/vol mixture of acetonitrile and 5 mmol/L phosphate buffer pH 7, marker thiourea. The line represents linear fit of experimental data.

considerable increase in flow velocity through the thermally initiated monolithic capillaries with the same level of charged moieties as the pore size increases from 250 nm to several micrometers [64]. A similar increase in flow velocity was observed for monoliths prepared by UV initiated polymerization [60,72]. Fig. 6.31 clearly demonstrates these effects. The range of pore sizes in question significantly exceeds the thickness of a few nanometers at which the electrical double layers would overlap for a system utilizing a mobile phase containing low molarity buffers [64]. Moreover, if it is assumed that the observed decrease in flow rate with decreasing mode pore diameter would simply result from the increasing percentage of pores within which overlap of the electric double layers occurs, then the flow velocity should reach a maximum value for those monoliths having sufficiently large pores, and remain constant thereafter, since the number of pores within which overlap of the electric double layer can occur decreases rapidly as the pore size increases. In practice however, this phenomenon is not observed. The fact that the overall flow velocity increases linearly over a broad range of pore sizes may also support the contention that this increase in flow rate is macroscopically related to a decrease in the resistance to flow through the channels. An additional effect may result from microscopic variations in the strength


Chapter 6

232

Fig. 6.32. Schematic of flow path in monolithic separation media with large (a) and small (2) pores.

of the electrical field in both the small and large pores. The effects of tortuosity and variations in the cross sectional area of a packed structure on the conductance and chromatographic performance of CEC capillaries packed with beads that have recently been discussed in the literature [75] may also play a role. It is likely, that the flow path through a monolith with large pores is rather straight and therefore, the trajectory is short. In contrast, this path may be longer in monoliths with small pores, since the molecules passing through the pores make many detours around a larger number of clusters of microglobules. As shown in Fig. 6.32, the analyte must traverse a longer distance that then results in longer retention times even if the overall electroosmotic velocity does not change. 6.10 OTHER APPLICATIONS OF POROUS POLYMER MONOLITHS IN CEC COLUMN TECHNOLOGY 6.10.1 Immobilization of beads in porous polymer monolith

A number of different approaches were used for monolithization of capillary column packed with silica beads [76-82]. Chirica and Remcho employed sol-gel

Monoliths

233

transition of silicate solution located in interstitial voids of a packed capillary column [83]. Since control of the solid phase formation to avoid cracks is very tricky [78,81], Chirica and Remcho used in situ polymerization of organic monomers [63]. Although the preparation of monoliths by polymerization of a mixture of monomers and porous particles was described in an earlier patent [84], this was the first application of this approach to prepare CEC columns. The process consists of a few steps: First, a very porous temporary frit is sintered at one end of the capillary and the tube slurry packed with porous 5 ~tm octadecyl silica beads. Using positive pressure, the packed capillary is filled with the polymerization mixture consisting of ethylene dimethacrylate, azobisisobutyronitrile, 1-propanol, 1,4-butanediol, and water. The last three compounds are inert solvents added to achieve the required porosity for the flow through. Optionally, an alkyl methacrylate, where the alkyl is methyl, ethyl, or butyl, and 2-acrylamido-2-methyl-1propanesulfonic acid were also added. After 48 h polymerization at a temperature of 60~ both ends of the capillary including that with the original frit were cut to obtain column with a desired length. SEM micrograph in Fig. 6.33 clearly shows the porous polymer within the voids between silica particles. Since the column is packed across its entire length, a piece of open capillary with a detection window was coupled this monolithic column. An addition of polymerizable sulfonic acid to polymerization mixture is a suitable tool for the control of electroosmotic flow. For example, only 0.1% of 2-acrylamido-2-methyl-1-propanesulfonic acid in the polymer increases electroosmotic flow velocity by 10-15% thus accelerating the separations. Despite numerous efforts, use of polymerization mixtures with only ethylene dimethacrylate as a monomer led to immobilized columns with dramatically decreased retentions. Since these mixtures are likely to produce porous polymers with very small pores and the polymer fills also the native pores of silica beds, the access of analytes to the C18 functionalities is restricted. Substitution of approximately half of ethylene dimethacrylate in the mixture with modestly hydrophobic methyl methacrylate returns the retention characteristics back to those of the original C18 silica. However, peak shapes featuring severe tailing were monitored for highly hydrophobic polyaromatic hydrocarbons. Using butyl methacrylate as a comonomer eventually alleviated this problem. As has been shown elsewhere [66], copolymers of ethylene dimethacrylate and butyl methacrylate exhibit hydrophobicity index similar to that of C 18 silica and this match of polarities is favorable for separations in the reversed phase mode. Fig. 6.34 demonstrates the performance of a column prepared using this technology for the separation of a series of drugs.


234

Chapter 6

Fig. 6.33. Scanning electron micrograph of a capillary column packed with 5 pm ODS silica beads and entrapped in porous poly(methyl methacrylate-co-ethylene dimethacrylate-co-2acrylamido-2-methyl-l-propanesulfonic acid). (Reprinted with permission from [63]. Copyright 2000 American Chemical Society).

6.10.2 Monolithic frits

In addition to utilization of monoliths as a column material, two reports describing respectively silicate and synthetic organic polymer based monolithic frits were published recently [85,86]. The conventional method of frit fabrication for a particle packed column usually involves thermal sintering of a section of the packing material, such as bare or octadecyl silica, using a heating device. This approach has several weaknesses such as the lack of control of the temperature and porous properties of the flit that decreases reproducibly of the fabrication process. In contrast, photoinitiated free radical polymerization of glycidyl methacrylate and trimethylolpropane trimethacrylate in the presence of porogenic solvent affords a monolithic plug within the column that serves as a frit. This procedure represents a simple approach to reproducible fabrication of frits even in capillaries with large inner diameters.

23 5

Monoliths

;k- 254 nm

e-,

L

,_____.

~ 220 nm

6

< 2 3

0

I 2

I 4

7

I 6

I 8

min

Fig. 6.34. CEC separation of drugs using a capillary column packed with 5 mm ODS silica beads and entrapped in porous poly(butyl methacrylate-co-ethylene dimethacrylate-co-2acrylamido-2-methyl-l-propanesulfonic acid). (Reprinted with permission from [63]. Copyright 2000 American Chemical Society). Conditions: column 26 cm (active length 17 crn) x 75 p,m i.d, mobile phase 70:30 acetonitrile-10 mmol/L acetate buffer solution pH 3, 20 kV, detection at 2 different wavelengths shown.

The initiation by UV irradiation facilitated localization of the polymer plug in the capillary column. An outlet frit was prepared first by introducing the monomer mixture into the capillary from which about 2 mm section of the polyimide coating was removed and its ends were sealed. The capillary was then masked by aluminum foil, leaving 1 mm of the section without coating exposed to the UV light. The polymerization process was finished in about 1 h, the unreacted monomer solution flushed from the tube, and the column packed with 1.5 lam octadecyl silica beads. Similar procedure was employed to form the inlet frit. Although no specific functionalization of the inner capillary surface preceding the polymerization was carried out, the outlet frit easily withstand a short exposure of a pressure as high as 400 MPa used during the column packing. This demonstrates that the 1 mm thick monolithic frit is strongly bound to the inner surface of the bare capillary wall. Fig. 6.35 shows the inlet frit structure with embedded 1.5 gm beads. This micrograph also suggests that the photopolymer does not exhibit the microglobular morphology typical of standard macroporous polymers (vide supra) if formed in the presence of the silica beads. The silica particles are entrapped in the pores and hold within their domains.


236

Chapter 6

Fig. 6.35. SEM micrograph of a cross section of a photopolymerized 1 mm long inlet frit with embedded 1.5 ~tm ODS beads within a 75 gm i.d. capillary. (Reprinted with permission from [86]. Copyright 2000 American Chemical Society).

In contrast to some other procedures, the UV photoinitiated polymerization does not require elevated temperature for the reaction to be completed. Therefore, the mobile phase used for packing remains in both the outlet frit and the packing during polymerization of the inlet frit. Consequently, the conditioning time for the column prior to its use is shortened significantly. No bubble formation was observed while using packed capillary columns with photopolymer frits. A systematic study of the run-to-run reproducibility for the analysis of a mixture of neutral molecules consisting of thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene was carried out over a period of 3 days. Figure 6.36 shows electrochromatograms of runs 10, 30, and 50. There is almost no variation in retention times of all test compounds. The relative standard deviations for retention factor, the efficiency, and the resolution, averaged over 60 runs, are 3.5%, 3.3%, and 5.5%, respectively. The integrity of the packed column remained unchanged, and the column

237

Monoliths

Run 10

r Run 30

r

r

~J r

Run 50

/x..__ I 2

,

I 4

I

[

6

I

I 8 min

Fig. 6.36. Selected representative CEC electrochromatograms from 60 runs using column with monolithic flits. (Reprinted with permission from [86]. Copyright 2000 American Chemical Society). Conditions: mobile phase 40% (v/v) 5 mM phosphate buffer with addition of 2 mmol/L SDS (pH 7.0) and 60% (v/v) acetonitrile. Peaks: thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene (order of elution).

is assumed to be useful for a much longer period of time. The column efficiencies calculated from peaks of thiourea, benzyl alcohol, benzaldehyde, and 2-methylnaphthalene shown in Figure 6.36, are 200 000, 160 000, 60 000, and 20 000 plates/m, respectively. 6.11 ACKNOWLEDGMENT I would like to thank my friend and colleague at the University of California at Berkeley, Professor Jean M. J. Fr6chet for his endless support and helpful advice, and co-workers Dr. Cong Yu, Dr. Michael L~immerhofer, Dr. Eric C. Peters, Dr. Miroslav Petro, Dr. David S3~kora for their most valuable contributions to the research and


238

Chapter 6

development of monolithic CEC columns that are listed in the references. Support of our research by grant of the National Institute of General Medical Sciences, National Institutes of Health (GM-48364) is gratefully acknowledged. Our work was also partly supported by the Division of Materials Sciences of the U.S. Department of Energy under Contract No. DE-AC03-76SF00098. 6.12 R E F E R E N C E S

1 J.H. Knox, J. Chromatogr. A, 680 (1994) 3. 2 M. Kubin, P. ~;pa6ek and R. Chrome~ek, Coll. Czechosl. Chem. Commun., 32 (1967) 3881. 3 W.D. Ross and R.T. Jefferson, J. Chromatogr. Sci., 8 (1970) 386. 4 F.D. Hileman, R.E. Sievers, G.G. Hess and W.D. Ross, Anal. Chem., 45 (1973) 1126. 5 H. Schnecks and O. Bieber, Chromatographia, 4 (1971) 109. 6 T.R. Lynn, D.R. Rushneck and A.R. Cooper, J. Chromatogr. Sci., 12 (1974) 76. 7 D. Josi6 and A. ~;trancar, Ind. Eng. Chem. Res., 38 (1999) 333. 8 K.H. Hamaker, S.L. Rau, R. Hendrickson, J. Liu, C.M. Ladish and M.R. Ladish, Ind. Eng. Chem. Res., 38 (1999) 865. 9 S. Hjert6n, Ind. Eng. Chem. Res., 38 (1999) 1205. 10 F. Svec and J.M.J. Fr6chet, Ind. Eng. Chem. Res., 36 (1999) 34. 11 S.M. Fields, Anal. Chem., 68 (1996) 2709. 12 H. Minakuchi, K. Nakanishi, N. Soga, N. Ishizuka and N. Tanaka, Anal. Chem., 68 (1996) 3498. 13 F. Svec, E.C. Peters, D. S3~kora, C. Yu and J.M.J. Fr6chet, HRC-J. High Resol. Chromatogr., 23 (2000) 3. 14 F. Svec, E.C. Peters, D. S~kora and J.M.J. Fr6chet, J. Chromatogr. A, 887 (2000) 3. 15 C. Fujimoto, Anal. Chem., 67 (1995) 2050. 16 C. Fujimoto, J. Kino and H. Sawada, J. Chromatogr. A, 716 (1995) 107. 17 N. Tanaka, K. Nakagawa, H. Iwasaki, K. Hosoya, K. Kimata, T. Araki and D.G. Patterson, J. Chromatogr. A, 781 (1997) 139. 18 B. Maichel, B. Poto6ek, B. Ga~, M. Chiari and E. Kenndler, Electrophoresis, 19 (1998) 2124. 19 B. Maichel, B. Poto6ek, B. Ga~ and E. Kenndler, J. Chromatogr. A, 853 (1999) 121. 20 B. Poto6ek, E. Chmela, B. Maichel, E. Tesa~ovfi, E. Kenndler and B. Ga~, Anal. Chem., 72 (2000) 74. 21 B. Maichel, B. Ga~ and E.Kenndler, Electrophoresis, 21 (2000) 1505. 22 M.R. Schure, R.E. Murphy, W.L. Klotz and W. Lau, Anal. Chem., 70 (1998) 4985. 23 N. Tanaka, T. Fukutome, K. Hosoya, K.Kimata and T.Araki, J. Chromatogr. A, 716 (1995) 57. 24 N. Tanaka, H. Iwasaki, T. Fukutome, K. Hosoya and T. Araki, HRC- J. High Resolut. Chromatogr., 20 (1997) 529. 25 B. Poto6ek, B. Maichel, B. Ga~, M. Chiari and E. Kenndler, J. Chromatogr. A, 798 (1998) 269. 26 Y. Baba and M. Tsuhako, YrAC-Trends Anal. Chem., 11 (1992) 280. 27 C. Fujimoto, Analusis, 26 (1998) M49.

Monoliths

23 9

28 S. Hjert6n, D. Eaker, K. Elenbring, C. Ericson, K. Kubo, J.L. Liao, C.M. Zeng, P.A. Lindstr6m, C. Lindh, A. Palm, T. Srichiayo, L. Valcheva and R. Zhang, Jpn. J. Electrophor., 39 (1995) 105. 29 C. Ericson, J.L. Liao, K. Nakazato and S. Hjert6n, J. Chromatogr. A, 767 (1997) 33. 30 J i . Liao, N. Chen, Ch. Ericson and S. Hjert6n, Anal. Chem., 68 (1996) 3468. 31 C. Ericson and S. Hjert6n, Anal. Chem., 71 (1999) 1621. 32 D. Hoegger and R. Freitag, HPCE 2000, 13th International Symposium on High Performance Capillary Electrophoresis and Related Microscale Techniques, Saarbri.icken, Germany, February 20-24, 2000, Abstracts p.230. 33 C. Ericson, J. Holm, T. Ericson and S. Hjert6n, Anal. Chem., 72 (2000) 81. 34 J. Seidl, J. Malinsk~,, K. Du~ek and W. Heitz, Adv. Polym. Sci., 5 (1967) 113. 35 A. Palm and M.V. Novotny, Anal. Chem., 69 (1997) 4499. 36 P.G. Righetti, J. Chromatogr. A, 698 (1995) 3. 37 A.H. Que, T. Konse, A.G. Baker and M. Novotny, Anal. Chem., 72 (2000) 2703. 38 A.H. Que, A. Palm, A.G. Baker and M. Novomy, J. Chromatogr. A, 887 (2000) 379. 39 N.W. Smith and M.B. Evans, Chromatographia, 41 (1995) 197. 40 G. Wulff, Angew. Chem.,Intl. Ed. Engl., 34 (1995) 1812-1832. 41 S. Nilsson, L. Schweitz and M. Petersson, Electrophoresis, 18 (1997) 884. 42 L. Schweitz, L.I. Andersson and S. Nilsson, J. Chromatogr. A, 792 (1997) 401. 43 L. Schweitz, L.I. Andersson and S. Nilsson, Anal. Chem., 69 (1997) 1179. 44 L. Schweitz, L.I. Andersson and S. Nilsson, J. Chromatogr. A, 817 (1998) 5. 45 L. Schweitz, L.I. Andersson and S. Nilsson, Chromatographia, 49 (1999) $93. 46 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hobo, Chromatographia, 43 (1996) 585. 47 J.M. Lin, T. Nakagama, K. Uchiyama and T. Hubo, Biomed. Chromatogr., 11 (1997) 298. 48 J.M. Lin, T. Nakagama, X.Z. Wu, K. Uchiyama and T. Hobo, Fresenius J. Anal. Chem., 357 (1997) 130. 49 I. Gusev, X. Huang and C. Horvfith, J. Chromatogr. A, 855 (1999) 273. 50 X. Huang, J. Zhang and C. Horvfith, J. Chromatogr. A, 858 (1999) 91. 51 B.H. Xiong, L.H. Zhang, Y.K. Zhang, H.F. Zou and J.D. Wang, HRC-J. High Resolut. Chromatogr., 23 (2000) 67. 52 F. Svec and J.M.J. Fr6chet, Anal. Chem., 54 (1992) 820. 53 F. Svec and J.M.J. Fr6chet, Science, 273 (1996) 205. 54 E.C. Peters, M. Petro, F. Svec and J.M.J. Fr6chet, Anal. Chem., 69 (1997) 3646. 55 C. Yu, F. Svec and J.M.J. Fr6chet, Electrophoresis, 21 (2000) 120. 56 M. L~immerhofer, E.C. Peters, C. Yu, F. Svec, J.M.J. Frdchet and W. Lindner, Anal. Chem., 72 (2000) 4623. 57 Q. Wang, F. ~;vec and J.M.J. Fr6chet, Anal. Chem., 65 (1993) 2243. 58 C. Viklund, F. Svec, J.M.J. Fr6chet and K. Irgum, Chem. Mater., 8 (1996) 744. 59 F. Svec and J.M.J. Frdchet, Macromolecules, 28 (1995) 7580. 60 M. L~immerhofer, F. Svec, J.M.J. Fr6chet and W. Lindner, Anal. Chem., 72 (2000) 4614. 61 F.E. Regnier, Nature, 350 (1991) 643. 62 D.K. Roper and E.N. Lightfoot, J. Chromatogr. A, 702 (1995) 3. 63 G.S. Chirica and V.T. Remcho, Anal. Chem., 72 (2000) 3605. 64 E.C. Peters, M. Petro, F. Svec and J.M.J. Fr6chet, Anal. Chem., 70 (1998) 2288.

240

Chapter 6

65 66 67 68 69 70 71

P.B. Wright, A.S. Lister and J.G. Dorsey, Anal. Chem., 69 (1997) 3251. E.C. Peters, M. Petro, F. ~;vec, and J.M.J. Fr6chet, Anal. Chem., 70 (1998) 2296. E. Venema, J.C. Kraak, H. Poppe and R.Tijssen, Chromatographia, 48 (1998) 347. E. Venema, J.C. Kraak, H. Poppe and R. Tijssen, J. Chromatogr. A, 837 (1999) 3. R. Stol, W.T. Kok and H. Poppe, J. Chromatogr. A, 853 (1999) 45. C. Fujimoto, Y. Fujise and E. Matsuzawa, Anal. Chem., 68 (1996) 2753. E.C. Peters, K. Lewandowski, M. Petro, F. ~;vec and J.M.J. Fr6chet, Anal. Commun., 35(1998) 83. C. Yu, F. Svec and J.M.J. Fr6chet, Electrophoresis, 21 (2000) 120. D. Li and V.T. Remcho, J. Microcol. Sep., 9 (1997) 389. J.H. Knox and I.H. Grant, Chromatographia, 32 (1991) 317. G. Choudhary and C. Horv~th, J. Chromatogr. A, 781 (1997) 161. M.M. Dittman, G.P. Rozing, G. Ross, T. Adam and K.K.Unger, J. Capil. Electroph., 5 (1997) 201. R. Asiaie, X. Huang, D. Faman and C. Horv~th, J. Chromatogr. A, 806 (1998) 251. M.T. Dulay, R.P. Kulkami and R.N. Zare, Anal. Chem., 70 (1998) 5103. Q.L. Tang, B.M. Xin and M.L. Lee, J. Chromatogr. A, 837 (1999) 35. S.A. Rodriguez and L.A. Colon, Anal. Chim. Acta, 397 (1999) 207. C.K. Ratnayake, C.S. Oh and M.P. Henry, HRC-J. High Resolut. Chromatogr., 23 (2000) 81. Q.L. Tang and M.L. Lee, HRC- J. High Resolut. Chromatogr., 23 (2000) 73. G. Chirica and V.T. Remcho, Electrophoresis, 20 (1999) 50. J.M.J. Fr6chet and F. Svec, US patent 5,334,310 (1994). Y. Chen, G. Gerhardt and R. Cassidy, Anal. Chem., 72 (2000) 610. J.R. Chen, M.T. Dulay, R.N. Zare, F. Svec and E.C. Peters, Anal. Chem., 72 (2000) 1224.

72 73 74 75 76 77 78 79 80 81 82 83 84 85 86

Chapter 7

Open Tubular Approaches to Capillary Electrochromatography r162

Joseph J. P E S E K

and M a r i a T. M A T Y S K A

Department of Chemistry, San Jose State University, One Washington Square, San Jose, CA 95192, USA

CONTENTS

7.1 7.2 7.3 7.4

7.5

7.6 7.7 7.8

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l etching process . . . . . . . . . . . . . . . . . . . . . . . . . C h e m i c a l modification process . . . . . . . . . . . . . . . . . . . . . . . Characterization o f etched, c h e m i c a l l y m o d i f i e d capillaries . . . . . . . . 7.4.1 Scanning electron m i c r o s c o p y . . . . . . . . . . . . . . . . . . . 7.4.2 A t o m i c force m i c r o s c o p y . . . . . . . . . . . . . . . . . . . . . 7.4.3 Diffuse reflectance infrared fourier transform spectroscopy . . . 7.4.4 Electron spectroscopy for chemical analysis . . . . . . . . . . . 7.4.5 E l e c t r o o s m o t i c flow m e a s u r e m e n t s . . . . . . . . . . . . . . . . Applications of OTCEC . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5.1 P h a r m a c e u t i c a l s and other small m o l e c u l e s . . . . . . . . . . . . 7.5.2 Peptides and proteins . . . . . . . . . . . . . . . . . . . . . . . 7.5.3 Chiral c o m p o u n d s . . . . . . . . . . . . . . . . . . . . . . . . . 7.5.4 Stability and reproducibility . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

242 244 245 247 249 250 253 255 256 257 258 260 263 264 266 267 268

242

Chapter 7

7.1 INTRODUCTION Applications and development of micro-separation techniques have expanded greatly in the last decade for a number of practical reasons. Among these are the reduced amount of sample needed, particularly important in biological and pharmaceutical applications, the lower volume of solvents used in the analysis, and the ease in which these methods can be interfaced to a mass spectrometer for a detector. Capillary electrochromatography (CEC) is one of the newest micro(~t)-separation formats available. In CEC there are two general approaches to this hybrid technique that combines features of both high performance liquid chromatography (HPLC) and capillary electrophoresis (CE): the packed column configuration that utilizes stationary phases similar to HPLC and the open tubular format where the stationary phase is immobilized on the capillary wall. This chapter focuses on the latter approach with an emphasis on open tubular columns that are fabricated by first etching the inner wall of the fused silica tube. A summary of the continuum of techniques that exist between ~t-HPLC and CE that can be used in the micro-column format are shown in Fig. 7.1. It can be seen that it is possible to make a smooth transition from Ia-HPLC to packed capillary CEC by adjusting the ratio of pressure-driven flow to electrically-driven flow. This adjustment results in an infinite number of hybrid techniques that can be generated from pure ~t-HPLC (no electric field component) to pure packed capillary CEC (no pressure component). The transition region between CE and packed capillary CEC is not so smooth. The bridge is provided by open tubular capillary electrochromatography (OTCEC). There are not, however, the infinite number of choices that can be generated as in the bridge between ~t-HPLC and packed capillary CEC. However, some variations exist by utilizing an etching process, which results in both an increase in surface area and radial extensions of various types and lengths from the surface, as well as the diameter of the capillary that can create a significant array of column formats. The figure also denotes on a relative scale the importance of the major separation mechanisms, solute/bonded phase interactions (as measured by the capacity factor, k') and electrophoretic mobility (laep), that exist in the continuum with k' being dominant at the ~t-HPLC end and ~tep being the only factor in CE. The wall of a fused silica capillary, containing silanol groups, has been identified as a potential site of interaction for solutes in electrophoretic measurements. In particular, adsorption of proteins, peptides and other basic compounds has been more specifically addressed [1-3]. In the ideal situation, the interaction is strictly chromatographic so that the process is reversible with the net result being a loss in the observed efficiency for CE [2,4]. A more serious problem can result when there is irreversible adsorption of the analyte on the capillary wall leading to a lack of repro-

243

Open Tubular C E C

~t-HPLC r

Voltage Assisted r p -HPLC

LAMINAR FLOW

~

~

Pressure Assisted CEC

~

~

~

~

Packed Capillary CEC

~

ELECTRICALL Y DRIVEN FLOW

O T C E C

r

CE

~- ~-- +- ~-- ~- Increasing influence of solute/bonded phase interactions (k') Increasing influence of electrophoretic mobility (~ep) --~ --~ -+ --~ --+ Fig. 7.1. Schematic that summarizes the relationship between the various micro-separation techniques. Line 1 lists the various micro-separation techniques. Below each method in line 2 is the type of flow or combination of flow ( ~ ) that exists for each technique while in line 3 is the relative contribution of solute/bonded phase interactions and electrophoretic mobility to the separation mechanism.

ducibility in crucial measurement parameters such as elution time, efficiency and even mobility. If the interaction is reversible, the specific solute/wall effect can be measured in well-designed capillary chromatographic experiments. Such measurements have been made in both the zonal [5,6] and frontal [7,8] chromatographic modes. The retention factor determined from these experiments can be used to determine the magnitude of the chromatographic interaction that appears in the efficiency term for CE [2,4,6]. The first effective OTCEC separations indicating chromatographic interactions were demonstrated a number of years ago by Tsuda and co-workers [9] using an octadecyl modified 30 pm i.d. capillary. A more definitive way of proving that chromatographic effects are possible in the open tubular format can be provided through the separation of optical isomers. A number of chiral selectors such as cyclodextrin and several cellulose derivatives that were bonded to the inner wall of fused silica capillaries resulted in the separation of enantiomers [ 10-15]. Since the two optical isomers have identical electrophoretic mobilities, separation can only be achieved through differences in solute/bonded phase interactions. It has also been demonstrated that molecularly imprinted polymers bonded to the inner wall of a fused silica capillary can be another approach for a stationary phase in OTCEC [ 16]. Each of these early studies showed that OTCEC was feasible at least for the limited number of samples and experimental conditions tested. Initially, further exploitation of the OTCEC technique was hampered by two fundamental problems that seriously limited its potential as a viable separation method: the low capacity of the column due to the small area available for bonding a stationary phase and the long distance that molecules would have to migrate to interact with the References pp. 268-270

244

Chapter 7

bonded moiety. The latter problem could be addressed by reducing the column i.d. but this is often an unsatisfactory solution because such a decrease further limits the sample size as well as the detection path length in optical systems and hence the detection limit of the analyte. The general approach described below attempts to overcome these two major problems though chemical etching of the inner wall of the capillary. This process produces structural changes within the capillary that are more favorable for OTCEC. First, the etching process increases the overall surface area of the capillary, perhaps by as much as 1000-fold [17], which allows more stationary phase to be attached to the wall thus increasing the capacity of the column. Second, the dissolution and redeposition of silica material during etching creates radial extensions from the wall that decrease the distance a solute must travel in order to interact with the stationary phase. This aspect is especially important for separations of biomolecules where the diffusion coefficient is much lower than for typical small organic molecules or inorganic ions. 7.2 CHEMICAL ETCHING PROCESS The etching process as described in this section was patterned after the original work of Onuska and co-workers for gas chromatography [17] but was modified to accommodate the smaller inner diameters needed for OTCEC [ 18]. The general procedure starts with an approximate 2 m section of bare capillary having a 375 ~tm o.d. and a 20 or 50 lam i.d. that is filled with concentrated HC1, sealed and heated overnight at 80~ The tube is then flushed successively with deionized water, acetone and diethyl ether. The tube is dried for 1 h under nitrogen flow at ambient temperature. The capillary is then filled with a 5% (w/v) solution of ammonium hydrogen fluoride in methanol and allowed to stand for 1 h. The methanol is then removed by nitrogen flow for 0.5 h. After the capillary is sealed at both ends, it is heated in a modified gas chromatography (GC) oven at temperatures between 300 to 400~ for a period of 3 to 4 h. The GC oven provides good temperature control, rapid heating and cooling, and easy accessibility for the lengths of capillary used in this procedure. The exact time and temperature chosen determines the surface morphology that is obtained on the inner wall [18,19]. The general trend observed is that for shorter etching periods (- 2-3 h) at lower temperatures (-300~

the inner wall consists of relatively long

spikes of silica material protruding from the surface. For longer etching periods (- 4h) at higher temperatures (-400~ the structure has fewer long extensions from the surface and becomes somewhat more regular either resembling sand dunes or a sponge-like porous configuration. More detailed studies by atomic force microscopy (AFM) and scanning electron microscopy (SEM) for characterizing etched and etched chemically modified capillary surfaces are described below. Estimates from the AFM

245

Open Tubular CEC

measurements indicate that the area of the inner wall is increased significantly (100 1000 fold), which should facilitate solute/bonded phase interactions after appropriate organic moieties are attached to this etched surface. Additional measurements of the surface area have been made by the BET (Brunauer-Emmet-Teller) nitrogen adsorption method. The data was obtained on 1 cm segments of etched 50 gm capillary with a total length of 2.6 m. The surface area measured was 1 x 10-1 m2/g. The same length of unetched capillary should have a surface area of approximately 8 x 10-5 m2/g. Therefore, the BET and AFM measurements are in good agreement showing an increase in surface area by about a factor of 1000. Additional useful surface morphologies may be possible through manipulation of the various experimental variables involved in the etching process, i.e. etching time, etching temperature, reagent concentration, capillary diameter and capillary configuration in the oven. 7.3 C H E M I C A L M O D I F I C A T I O N P R O C E S S After etching, bonding of an organic moiety to the etched inner wall for the stationary phase takes place using the silanization/hydrosilation process adapted for the capillary format [ 18-20]. This approach for attaching a variety of organic moieties to silica involves first silanizing the surface by reaction with triethoxysilane (TES) to create a hydride intermediate [21 ] as shown in equation 7.1. SILANIZATION OY H+

I

~ Si-OH + (OEt)sSi-H

~

~ ,g

(7.1)

Si-O-Si-H + nEtOH

I OY

First the capillary is treated with a pH 10 ammonia solution (6 mM) for 20 h at a flow rate of 0.1-0.2 mL/h. The capillary is then rinsed with deionized water followed by a wash with 0.1 M HC1 and then a second rinsing with water. The tube is next dried with nitrogen, filled with dioxane and then flushed with a 1.0 M TES solution in dioxane containing HC1 as a catalyst for 90 min at 90~

After the TES treatment, the

capillary was washed with tetrahydrofuran (THF) for 2 h and then with a 1:1 THFwater mixture for 2 h. The capillary is then dried with a flow of nitrogen gas for 0.5 h. The capillary, which now has a hydride surface, is then flushed with dry toluene in preparation for the second modification.


Chapter 7

246

The next step is the hydrosilation reaction that is used to attach an organic moiety [22,23]. An olefin/catalyst (usually hexachloroplatinic acid) solution (pure olefin or olefin dissolved in toulene) is heated to 60-70~

for a period of 1 h. The hydride

capillary filled with toluene is then flushed with the reacting solution for a period of 45 h at 100~

After completion of the hydrosilation reaction, the capillary is washed

successively with toluene and THF for 1 h. After the washing step, the capillary is then dried at 100~ under a nitrogen flow overnight. Normally hydrosilation utilizes as a terminal olefin for creating a Si-C bond between the silicon containing reactant and the organic compound [24] as shown in equation 7.2. HYDROSILATION

I

Si-H

cat.-- I~ S i - C H 2

+ R - C H m CH2

m CH2mR

(7.2)

c a t - catalyst, typically hexachloroplatinic acid (Speier's catalyst) However, it has already been demonstrated that other unsaturated groups (alkynes and cyano) as well as olefins in a nonterminal position can undergo hydrosilation on a silica hydride surface [25,26]. The cyano group is amenable to hydrosilation only in the absence of an olefin and with a free radical initiator as the catalyst giving two possible products as shown below [27].

/0 Ar m

~ N ~Si ~ 0 \

0

O\ / 0

/$i~ Ar

\

/ Si

/\ 0

0

247

Open Tubular CEC

We have also used free radical initiators and other transition metal complexes besides Speier's catalyst to attach some olefins to the silica hydride intermediate [26,28]. The silanization/hydrosilation method is a very versatile approach for attaching a wide variety of organic moieties to silica surfaces. This includes porous silica particles for HPLC and packed capillary electrochromatography, and to the inner walls of fused silica tubes for use in CE and OTCEC. Another advantage of the silanization/ hydrosilation method is that most of the readily accessible silanols are eliminated creating a surface that is particularly suited for the analysis of biomolecules and pharmaceuticals. Finally, when alkenes or alkynes are used in the hydrosilation reaction, bonding to the silica surface results in a highly stable Si-C linkage. The type of bonded phase and hence the specific solute/bonded phase interactions that can be exploited for any particular separation depends on the nature of the organic moiety attached to the surface. The types of groups bonded to the etched inner wall of fused silica capillaries have been hydrophobic (octadecyl, C18) [18-20,29,30], hydrophilic (diol) [30], chiral [31] and liquid crystals [32]. The structures of the molecules attached to the silica hydride surface are shown in Fig. 7.2. In some cases linkers are first bonded to the desired stationary phase moiety before hydrosilation. The versatility of the silanization/hydrosilation process that was proved for the synthesis of bonded phases for HPLC can be readily adapted to OTCEC. As will be shown below, the etching process does make some contribution to the overall chromatographic/electrophoretic behavior of the separation process in OTCEC using the etched chemically modified capillaries. A summary of the complete process for producing OTCEC columns by the two steps (etching and chemical modification) described above is shown in Fig. 7.3. 7.4

CHARACTERIZATION

OF

ETCHED,

CHEMICALLY

MODIFIED

CAPILLARIES

A number of different approaches have been developed for characterizing the etched capillaries. Just as the development of methods for characterizing chemically bonded stationary phases has been useful in understanding their performance, a similar strategy is essential for future development of these separation materials as well as the identification of appropriate applications for OTCEC. In contrast to the porous particles typically used in HPLC where the chemically modified surface is readily accessible to various spectroscopic techniques [33,34], probing the inner wall of a fused silica capillary is not a straightforward process. In most cases, it is necessary to open the capillary so that the inner surface is exposed. This is not a simple procedure. It usually involves carefully breaking the capillary along the longitudinal axis and then examining the fragments under an optical microscope in order to select the most


248

Chapter 7

HYDROPHOBIC: Octadecene CH2~--CH- CH2-- (CH2)I4~CH3

HYDROPHILIC" Diol OH OH

I

I

CH2~-CH- (CH2)4--CH--CH2

CHIRAL: R(+)-l,-(a - naphtyl)ethylamine 0 CH3 Cyclodextrin

..H O"

OH

O

I

II

1~= ~ C H 2 - - C H - C H 2 ~ O ~ C ~ C - - C H 2

I

CH3

OR' LIQUID CRYSTALS: Cholesteryl l O-Undecanoate

C ~ : ~- - 'H O

\

CH3

C H 2 : C H - CH2-- (CH2)6 ~ C H 2 - - C -O

I~I

H 4-cyano-4Ln-pentoxybiphenyl (M) C 5H l 1 - - O ~ ~ ~ ~ -

\~-~/

~

CN

Fig. 7.2. Structures of molecules bonded to the etched inner walls of fused capillaries by the silanization/hydrosilation process. In some cases a linker molecule has been attached to the primary separation moiety in order to facilitate bonding to the hydride surface.

249

Open Tubular CEC

CEC Capillary Derivatization . . . . . . . . . . . . . . . :........

--

-

-

native

~

ylVv

,v

~,

etched

~

yIV"

v-

Vv

hydrided

~

IFlV"

,v

V'

alkylated

I

50

~.tm

20 pm

or

NH4HF2

silanization: SiH(OEt)3

hydrosilation (alkylation): RCH=CH2 + catalyst

Fig. 7.3. Schematic representation of the processes for producing etched, chemically modified capillaries for OTCEC.

suitable samples for a particular characterization technique. Useful fundamental information about the properties of the etched chemically modified surface has been obtained from the five methods described below.

7.4.1 Scanning Electron Microscopy (SEM) SEM provides a picture of the surface morphology that exists on the inner wall of the capillary. This image is particularly useful for comparing the original smooth wall to the greatly roughened surface after etching as well as for comparing the effects of various etching conditions with respect to the type of new three-dimensional structures formed. The sample used is obtained as described above since the inner surface must be accessible to the probing electron beam. After a suitable sample is identified, the surface is gold plated by a sputtering process in order to make the capillary conductive. A typical SEM image showing the inner wall of a fused silica capillary after etching is shown in Fig. 7.4. A variety of 20 and 50 pm etched capillaries have been examined by SEM with respect to trying to correlate the surface morphology obtained as a function of the etching conditions [18,19]. The structures as described above in the section on the ammonium hydrogen difluoride process indicate that for short etching periods, at least two hours and at a temperature of 300~

the surface

appears to have the greatest amount of roughness and longer radial extensions from the surface. Using longer etching times and/or higher temperatures generally results in a diminishing of these features as the original extensions from the inner wall "melt away to a less tortuous morphology. At best, SEM provides a good visual image of the surface but only a semi-quantitative description of the overall roughness that results from the etching process.


250

Chapter 7

Fig. 7.4. Scanning electromicrograph of the inner wall of an etched fused silica capillary.

7.4.2 Atomic Force Microscopy (AFM) AFM offers an alternative to SEM in providing a topological description of the etched, chemically modified surface. The two methods in combination provide complementary and confirming characterization of the inner wall that can be used to help understand the separation process in OTCEC using this format. The advantages of AFM over SEM include the ease of sample preparation because a conducting surface is not required, more precise determination of surface roughness from higher resolution in the z-direction (access to height information that is not available in SEM), and the ability to measure localized surface forces in order to identify possible solute/bonded phases interactions. The disadvantages of AFM for the characterization of OTCEC capillaries center on the difficulties in physically accessing regions of convoluted surfaces and uncertainties in rendering non-planar surfaces. The latter problem arises from contributions of the scanning piezo with respect to its movement over a curved surface. However, earlier AFM studies [35,36] have shown that some of the problems associated with non-planar and non-uniform surfaces of capillaries coated with polyacrylamide on the inner wall can be overcome. A comparison of SEM and AFM images of the same surface for the inner bore of native, etched, and etched chemically modified capillaries revealed a good visual

251

Open Tubular CEC

native

etched

hydride

chem-mod

Fig. 7.5. Atomic force microscopy (AFM) images in 3D recorded at two scan sizes of the four stages involved in the etching and chemical modification of capillaries for OTCEC.

correspondence between the electron and force microscopy data [37]. However, there are resolution differences between the two techniques. Subtle features revealed in AFM images require high magnification by SEM that limits the scan size to areas too small to observe the larger surface components. Some examples of the images obtained by AFM of each stage in the production of etched chemically modified capillaries are shown in Fig. 7.5. Two different scan sizes are displayed for the native, etched, etched-hydride and etched-chemically modified capillaries. The higher magnification (smaller scan size) allows for the examination of the more subtle features on each surface. Both scan sizes reveal that the bare capillary is relatively flat without any prominent features while the etched surface displays significant roughening with fairly sharp extensions. The roughness seems to increase further upon silanization to create a hydride layer. The rugged nature of the hydride surface may be due to the fact that more than a single layer of silane may have been deposited during this process. The micrographs of the alkylated surface reveal that considerable filling in and smoothing occurs during the hydrosilation process. Both AFM and SEM can easily detect the differences in surface morphology present in the four types of capillaries that are relevant to this approach to OTCEC. Besides images of the surface, AFM provides additional information that makes it unique as a surface characterization method for the etched, chemically modified capillaries. These measurement features include root means square (RMS) roughness,


252

Chapter 7

surface area and surface-tip forces of attraction. When these AFM measurements are made on the four types of capillaries involved in the fabrication of etched, chemically modified columns for OTCEC, several crucial observations can be made about these unique separation materials. The ordering of the various surfaces for the RMS roughness measurements is native < etched ~ alkylated < hydride. The data is presented graphically in Fig. 7.6. For the surface area determinations the order is as follows:

NATIVE

40

,,,

ETCHED 30

, ,

20

0

40 HYDRIDED 30

. . . . . . .

20 10

li.....

0

.

m i

ALKYLATED

,,,

I

i

ml l,m,,_. 5

.

15

25

35

45

55

RMS ROUGHNESS, nm

65

75

Fig. 7.6. RMS roughness measurements as determined by AFM for the four types of capillaries involved in the etching and chemical modification processes.

Open Tubular CEC

253

native < etched < alkylated < hydride. A close agreement between these two parameters might be expected when comparing surfaces that change as a result of etching and chemical modification since an increase in the surface area should closely parallel an increase in the roughness. The actual order is interesting and might be explained by the following reasoning. The etched surface is greater than the native surface as expected from the etching process. The increase in going from the etched to the hydride surface may be the result of some polymerization of the silanization reagent, TES, giving rise to irregular growths. The smoothing of the hydride surface upon alkylation may be due to the infilling of the pits by alkyl groups attached during hydrosilation [38]. The relative magnitudes of the surface-tip forces of attraction are in the following order: native ~ alkylated ~ hydride I < etched < hydride II. The hydride surfaces have two distinct populations of forces so they are separated in the ordering scheme shown above. The ordering of the surface-tip forces can be qualitatively explained by the relative reactivities of the four surfaces. The native surface has been exposed to atmospheric condition over a long period and thus has fairly low surface energy. The etched surface with increased surface area is significantly more active and hence possesses higher energy. The hydride I surface is similar to native silica, passivated, while hydride II is activated like the etched surface. The alkylated surface has low surface energy similar to most hydrocarbon materials. The increased surface area that is covered by a substantial layer of organic material in the etched chemically modified capillaries can be correlated to the electrokinetic chromatographic behavior of these columns. More specific examples are given in the sections below but some general observations can be made with respect to the AFM data. The elution time for all compounds on the etched chemically modified capillary is significantly longer than on a chemically modified capillary that has not been etched indicating the presence of solute/bonded phase interactions. The peak width of solutes on the etched chemically modified capillary is greater than that for the same solutes that migrate through a native capillary indicating the presence of mass transfer effects. The elution times of solutes are longer as the diameter of the capillary diminishes because of larger k' values that result from the shorter distance a solute must diffuse in order to interact with the bonded moiety on the etched surface [39].

7.4.3 Diffuse Reflectance Infrared Fourier Transform (DRIFT) Spectroscopy Another limitation to the spectroscopic characterization of the inner wall of the etched fused silica capillary is the relatively small surface area. While the etching process is designed to increase the surface significantly with respect to the native material, the area is still small ( 495 nm). Reproduced with permission from Palm and Novotny [36]. Copyright 1997 American Chemical Society.

10.6 FATTY ACIDS

Fatty acids have so far been analysed by CEC either as the free acids or as phenacyl- or methyl esters. Aqueous acetonitrile (50 mM) at pH 6 (9:1 v/v) was shown to be the optimal mobile phase [ 191 ]. It is generally known that in reversedphase chromatography free fatty acids and fatty acid methyl esters separate according to the partition number, which is defined as the carbon number minus twice the number of double bonds. A double bond reduces the retention time by the equivalent

Applications

351

25

Glc7 Glc$ I Glc6

o,~ II 1'6

1'8

2'0 2'2 2'4 2~6 2'8 Time (min)

i

30

Fig. 10.8. Isocratic electrochromatography of the oligosaccharide ladder in a capillary filled with a macroporous polyacrylamide/poly(ethylene glycol) matrix, derivatized with a C4 ligand (15%) and containing vinylsulfonic acid (10%). Conditions: capillary, 500 mm (400 mm effective length) x 100 lam i.d.; mobile phase, acetic acid 1:1000 containing 5% (v/v) acetonitrile; field strength, 600 V/cm; injection 5 s (100 V/cm); detection, LIF (helium-cadmium laser, excitation at 325 nm, emission at > 495 nm). Reproduced with permission from Palm and Novotny [36]. Copyright 1997 American Chemical Society.

of two carbons (pairs corresponding to this rule are called "critical pairs"). With acetonitrile-50 mmol/1 MES [2-(N-morpholino)ethanesulfonic acid], pH 6 (9:1), and a run-voltage of 30 kV per capillary (30 or 40 cm long, 100 lam i.d.), good results can also be obtained. Fatty acids and fatty acid phenacyl esters originating from vegetable oils and margarines were subjected to capillary electrochromatography by Dermaux et al. [51 ]. Using the Hewlett Packard system for capillary electrophoresis pressurized from both sides at 10 bar, and fused silica capillary columns 35/25 and 50/40 cm long (100 lam i.d.) (the latter number specifies the active length of the packed bed), slurry-packed with 3 lam Hypersil ODS, the results shown in Fig. 10.10 were obtained. This Figure shows not only the comparison of results obtained using CEC with two different lengths of the packed bed, but also a comparison with micro reversed-phase chromatography (position A). Fatty acid phenacyl esters were run in acetonitrile-50 mM MES pH 6 (90:10), at 30 kV and 20~

the injection was electrokinetic. A comparison

of the separations obtained with derivatized and underivatized fatty acids is given in Fig. 10.11. As one would expect, it is also possible to separate at least some positional and geometrical isomers (Fig. 10.12).


352

Chapter 10

!

0.00! f

a.

0.006 0.004

o

[

5

5,6

4

0

~

10 3

----B 4

6

A -2

0

i

1

[

i

i

i

5

10

15

20

25

30

Time / rain Fig. 10.24. Separation of benzodiazepines by CEC. Column, 200 mm (effective length) x 100 pm i.d. packed with cholesteryl-bonded silica. Mobile phases: (A), acetonitrile-5 mM Tris-HC1 buffer pH 7.7 (35:75 v/v); (B), acetonitrile-5 mM Tris-HC1 buffer pH 73 (35:65 v/v). Applied voltage 300 V/cm. Reproduced with permission from Jinno et al. [ 176].

similar to that in the isocratic run, and it was proposed that during this period the second buffer did not yet penetrate the capillary column while the remaining two solutes apparently experienced the gradient effects and gave very sharp peaks.

10.13 ANTIBIOTICS Antibiotics represent a chemically quite heterogenous category of compounds, for which the C 18 phases appear sorbents of the first choice. A group of these compounds (tetracyclines) was successfully separated by open tubular capillary electrochromatography using C18 modified capillaries. The results reported to be obtained in this system were better than those obtained with either monolithic columns or open diol columns [123] (Fig. 10.26).

369

Applications

H

mAU

0

.

SCH=

13

80

o

14

(111

15

1121

16 9 SCH3

o~

o~~

~

12

1131

1141

o~

~

o~ t ~ ,

1151 . . . . . . . . . . . . . . . . . . . . . . .

.

.

.

.

,,j

.

.

,

.

.

.

.

i

,,...

,.

L

.............

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

---~

.

.

(111

l,~

. . . .

.

.

.

: :~

--

.

.

-

.-

.

!

Fig. 10.25. Separation of tipredane (compound 14) from its five related substances (compounds 11-13, 15, 16) by CEC. Column: 250 mm x 50 ~tm i.d. packed with 3 ~tm Spherisorb ODS-1; mobile phase, acetonitrile-50 mM Tris buffer, pH 7.8, 80:20 (v/v); applied voltage, 15 kV; temperature, 15~ electrokinetic injection, 5 kV/15s. From Euerby et al. [63], @Journal of Microcolumn Separations, 1997. Reproduced with permission of John Wiley & Sons, Inc.

10.14 PESTICIDES So far, capillary electrochromatography has found only a few applications to pesticides. Separation of cinosulfuron and its by-products can serve as a good example. The separation was done with a C-8 non-endcapped 3 ~tm packing in a capillary whose dimensions were 33.5 cm (25.0 cm to the detector) x 100 ~tm internal diameter. A typical separation is presented in Fig. 10.27 [ 128]. 10.15 AVAILABLE A P P L I C A T I O N S (summarizing Table) It is evident that the different applications of electrochromatographic procedures are growing every day. Because some of the procedures represent only small variations of published procedures already applied to another category of compounds, we have decided to supply the basic information available for the various categories of compounds discussed briefly in the preceding sections. Although we have attempted to make this Table as complete as possible, we cannot guarantee that all application papers are included, for a number of reasons. Nevertheless we believe that this rather extensive Table can serve as a source of information for readers and can help newReferences pp. 413-419

3 70

Chapter 10

mAU 29,5

A

2r

29

B

28.5

28.5~

2~ 27.~

26

3,b

27.5 27.

27

26.5-

26.5

26-

26

25.5

25.5

25

25

24.5

24.5

3,b

II

2423.523. . . .

3.~o . . . .

41o ....

~.~o

slo

TIME (mln)

C

28.52R

22.5t~

. . . . mAU

a71 a~. ]~

27.5 27

II

3,b

26.5

33. 32. al,

26

30

25.5

29.

~.~o''' '6.bo''' ~.go''' ?,oE~ ' '7.~o' TIME (rain)

3 /

, 9

II

D b

/ Ill 2

28.

25

27. 24.5

2r,

24 23.5

TIME (mini

~.bo' ' ' i3.bo'' ~.bo''' ioJo~" itJod ' ' i2:oo TIME (mini

Fig. 10.26. Separation of some likely components found in commercial tetracycline. (A), Bare capillary with buffer pH 3.0 (30 mM citric acid and 24.5 mM 13-alanine) 500 mm x 50 pm i.d.; 30 kV, electrokinetic injection for 5 s at 10 kV; (B), bare capillary, same as for A but with buffer-methanol (60:40 v/v); (C), Hydride (etched) capillary 250 mm x 50 pm i.d., other conditions same as for B; (D), C18 etched capillary, conditions same as for C. Peak identification: 2, tetracycline; 3, chlorotetracycline; a, 4-epitetracycline; b, anhydrotetracycline; e, 4-epianhydrotetracycline. Reproduced with permission from Pesek and Matyska [123].

comers to appreciate the categories (sets) of compounds for which capillary electrochromatography can be exploited.

Applications

3 71

N OCH3

UV-absorbance at 209 nm [mAU]

Cinosulfuron ~y- SO2-NH-CO-NH-~N 0CH2CH20CH3

N OCH3

120 100 1234

80-

6

604020_ I

I

6 1 Methanol 3 2 Thiourea

H3CO N

N~ ~ NH2 H3C0/-N 0,2%

7 4

0CHzCHzCCH3

I

I

N 0CH3 N\NH 2

0,2%

I

8 9 Time [m]

10 5

I

I

11

12 6

0CH2CH2CCH3

0,8%

0CH3

0,7%

Fig. 10.27. CEC separation of Cinosulfuron and by-products. Column: 335 (250) mm x 100 pm i.d. packed with Synchropak non-endcapped C8, 3 pm, Conditions: mobile phase, acetonitrile-20 mM sodium dihydrogenphosphate, pH 4.0 (60:40 v/v); separation voltage, 15 kV; injection, 5 kV, 3s; applied pressure, 8 bar on both sides; sample, 2.4 mM cinosulfuron dissolved in mobile phase. Reproduced with permission from Rapp et al. [ 128].


TABLE 10.1

taO "--1 bO

APPLICATIONS (compositions of mobile phases are described using volume proportions, i.e., v/v)

Compound

Stationary phase

Mobile phase

Capillary dimensions

Note

Ref.

Acetonitrile-water (40:60) pH 9.1

500 mm x 50 ~tm i.d.

420 mm to the detector, 6 25~

Hydrocarbons and fullerenes Alkylbenzenes (toluene, ethyl- 40% Ethylacrylate, 50% benzene, propylbenzene, methacrylic acid, 10% lauryl butylbenzene) methacrylate (custom-made polymer), 4 wt.% polymer concentration Polyaromatic hydrocarbons (benzene, toluene, naphthalene, acenaphthylene, fluorene, anthracene, 1,2-benzanthracene, phenol, acetone)

ODS Hypersil, 5 gm

Gradient acetonitrile-water (from 60:40 to 90:10)

500 mm x 75 ~tm i.d.

200 mm packed length, flow-injection analysis

7

Polyaromatic hydrocarbons

40% Ethylacrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 3.99-4.01 wt.% polymer concentration

Acetonitrile-buffer (40:60); pH 9.2 (10 mM sodium borate) or pH 11.5 (10 mM phosphate)

300 mm x 50 ~tm i.d.

250 mm to the detector

6

Polyaromatic hydrocarbons and parabenes (ethyl-hexyl parabene, naphthalene, fluorene, phenanthrene, fluoranthene, pentylbenzene)

Excil C 18

Acetonitrile-Tris HC1 buffer pH 8.0 (80:20)

380 mm x 100 p,m i.d.

85 mm packed length

8

"x

"~ .~ ~, & ~

Polyaromatic hydrocarbons Micra ODS-1, 1.5 gm or (benzene, naphthalene, Micra ODS-2, 3 gm acenaphthylene, fluorene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene, benz[a]anthracene, chrysene, benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[a]pyrene, dibenz[a,h]anthracene, indeno [ 1,2,3-cd]pyrene, benzo[ghi]perylene)

Acetonitrile-10 mM 440 mm x 50 ~m i.d. ammonium acetate, pH 6.5 (70:30) or acetonitrile-50 mM Tris buffer, pH 8.1 (80:20)

Polyaromatic hydrocarbons Spherisorb ODS-1, 3 gm or (benzene, naphthalene, Spherisorb ODS 2, 3 gm acenaphthylene, fluorene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene, benz[a]anthracene, chrysene, benzo[b]fluoranthene, benzo[k]fluoranthene, benzo[a]pyrene, dibenz[a, h] anthracene, indeno [ 1,2,3-cd]pyrene,

Acetonitrile-50 mM Tris buffer, pH 8.1 (80:20)

430 mm x 50 ~tm i.d. 350 or 410 mm packed or 530 mm x 75 p.m i.d. length

Acetonitrile--0.1 M acetate buffer, pH 3.0 (80"20)

250 or 170 mm (non-entrapped or entrapped), 75 gm i.d.

340 mm packed length, 9 voltage programming tested

r,,~~

r,,~~

t~

9

benzo[ghi]perylene) Polyaromatic hydrocarbons Nucleosil ODS, 5 gm (naphthalene, fluorene, phenanthrene, anthracene), acetone

250 and 170 mm are 10 effective column lengths

ta~

ta9 4~

TABLE 10.1 (continued)

Compound

Stationary phase

Mobile phase


Note

Ref.

Polyaromatic hydrocarbons (naphthalene, fluorene, phenanthrene, pyrene, benz[a]anthracene

Linear polymer coated capillary [poly(N-tert.-butyl acrylamide-co-2-acrylamido2-methyl- 1-propanesulfonic acid]


600 mm x 25 ~tm i.d.

450 mm effective column length

11

Polyaromatic hydrocarbons (benzene, toluene, ethylbenzene, propylbenzene, butylbenzene, pentylbenzene)

Separation segment: ODS, 5 pm (200 mm), accelerating segment: bare silica (0-280 mm)

Acetonitrile-1.25 mM monobasic sodium phosphate pH 6.0 (75:25)

100 lam i.d., for length Two types of packing see stationary phase

12

Polyaromatic hydrocarbons (sixteen US EPA PAH)

SynChrom ODS (3 pm, 90%) Acetonitrile-4 mMborate + Spherisorb SW (80:20) (1 lam, 10%)

330 mm x 75 gm i.d.

13

Polyaromatic hydrocarbons (sixteen US EPA PAH)

Nonporous ODS (90%) + 1 ~tm silica gel (10%)

Acetonitrile-2 mMTris, pH 9 200 (300) mm x 100 (65:35) pm i.d.

14

Fullerenes C60, C70

Vydac C18 (3 gm)

Acetonitrile-tetrahydrofuran (1"1)

400 mm x 5 gm i.d.

15

Alkylbenzenes

Custom made styrenedivinylbenzene polymer 3 and 6 lam

Acetonitrile-water (80"20)

300 mm x 100 pm i.d.

16 c~

~ ~ .~ ~" tao ~.

17

Polyaromatic hydrocarbons Nucleosi14000-7 C18, (naphthalene, fluorene, anthra- Nucleosil 1000-7 C 18, cene, pyrene), acetone Nucleosil 500-7 C 18

Acetonitrile-water (80:20) containing 10 mM borate pH 8.3

720 mm x 100 pm i.d.

Polyaromatic hydrocarbons Modified C18 open tubular (naphthalene, acenaphthylene, column acenaphthene, fluorene, phenanthrene, anthracene, fluoranthene, pyrene)

Methanol-1 mM phosphate buffer, pH 7.0 (65:35)

400 mm x 9.6 pm i.d.

Aromatic compounds (benzene, 1,2,3-trimethylbenzene, nitrobenzene, etc.)

Spherisorb ODS, 3 pm

Isocratic elution: acetonitrile- 435 mm x 75 gm i.d. 4 mMTris, pH 9.2 (60:40); gradient elution: acetonitrile4 mMTris, pH 9.2 (60:40)

158 mm packed length

19

Aromatic compounds (alkyl benzenes)

Spherisorb ODS-2, 3 pm

Acetonitrile-4 mMTris, pH 9.2 (80:20)

270 mm x 75 pm i.d.

66 mm packed length, high speed CEC

20

Aromatic compounds (alkyl benzenes)

Non-porous ODS, 2 pm


270 mm x 75 gm i.d.

63 mm packed length, high speed CEC

20

Aromatic hydrocarbons (naphthalene, 2-methylnaphthalene, fluorene, phenanthrene, anthracene)

2-Hydroxymethacrylatepiperazine copolymer with C 18 ligands and immobilized dextran sulfate

62% Acetonitrile in 5 mM borate, pH 8.7

550 mm x 25 pm i.d.

525 mm active length

21

Aromatic hydrocarbons (2-methylnaphthalene, fluorene, phenanthrene, anthracene)

2-Hydroxymethacrylatepiperazine copolymer with C 18 ligands

50% Acetonitrile in 4 mM sodium phosphate, pH 7.4 (60%), acetonitrile-buffer (60:40) or gradient from 60:40 to 70:30 acetonitrilebuffer

200 mm • 75 9m i.d.

160 mm active length; comparison of isocratic and gradient elution

22

r.,~~

300 mm to the detector, 18 optimization of mobile phase, open tubular electrochromatography

r~

taO

L~


Compound

Stationary phase

0.3 wt.% 2-acrylamido-2Benzene derivatives (benzyl methyl- 1-propan sulfonic alcohol, benzaldehyde, benacid, pore size 465 nm zene, toluene, ethylbenzene, propylbenzene, amylbenzene), thiourea Polystyrene standards (987.103 rel. mol. mass) and toluene

Butyl methacrylate-ethylene dimethacrylate copolymer containing 0.3 wt.% 2-acrylamido-2-methyl- 1-propanesulfonic acid

Note

Ref.

Mobile phase


Acetonitrile-4 mM phosphate buffer, pH 7 (80:20)

250 mm x 100 ~tm i.d. (250 mm active length)

23

Tetrahydrofuran containing 2 vol.% of water

300 m m x 100 ~tm i.d. (300 mm active length)

24

CEC Hypersil C 18, 3 gm Polyaromatic hydrocarbons and parabenes (thiourea, ethyl-hexylparabene, naphthalene, fluorene, phenanthrene, anthracene, fluoranthene)

450 mm x 100 gm i.d. Acetonitrile-water-25 mM Tris.HC1, pH 8.0 (gradient from 20 to 80% of acetonitrile)

250 mm to the detector 25

Hypersil C 18, 3 ~tm Aromatic hydrocarbons and other compounds (thiourea, benzyl alcohol, methyl benzoate, toluene, benzophenone, naphthalene, 1,4-dichlorobenzene, phenothiazine, biphenyl, 1,3,5-trichlorobenzene, 1,2,4,5-tetrachlorobenzene)


Packed length 250 mm, 26 separations at 60 ~ and 20~ compared (inclusive temperature gradient)

335 mm x 100 ~tm i.d.

Aromatic hydrocarbons (naphthalene, phenanthrene, pyrene) 4~

Tetraethoxysilane-noctyltriethoxysilane hybrid gels

Aromatic hydrocarbons, Sol-gel bonded 3 gm benzyl alcohol, benzaldehyde, ODS/SCX, 80 A pores benzophenone, biphenyl, benzene, toluene, ethyl- and butylbenzene

Acetonitrile-water (1" 1)

470 mm x 50 ~tm i.d.


27

Acetonitrile-water (80:20) containing 1.5 mM phosphate buffer, pH 3.0

340 mm x 75 lam i.d.


28

Acetonitrile-25 mM morpholinoethanesulfonic acid (95:5), pH 6.2

300 mm x 75 gm i.d.

1O0 mm packed length

29

Polynuclear aromatic hydrocarbons (naphthalene, phenanthrene, pyrene), thiourea

Particle-loaded (3 gm, C18) monolithic sol-gel column

Polyaromatic hydrocarbons (naphthalene, fluoranthene), thiourea

BDS-ODS Hypersil, ODS Hypersil, Spherisorb ODS-2, Spherisorb ODS- 1, CEC Hypersil C 18

Polyaromatics (ethyl-hexylparabenes, naphthalene, fluorene, phenanthrene, anthracene, fluoranthene), thiourea

Hypersil ODS, 3 gm

Acetonitrile-25 mM Tris.HC1 buffer, pH 8.0 (80:20)

335 mm x 100 (75) gm i.d.

Polyaromatics (naphthalene, acenaphthene, phenanthrene, anthracene, fluoranthene, pyrene)

Capillary had porous silica layer (0.70 ~tm) with chemically bonded C 18

Acetonitrile-1 mM phosphate buffer, pH 7.0 (50:50)

400 mm x 9.60 gm i.d. 300 mm to the detector, 32 open tubular CEC

Comparison of different 30 C 18 stationary phases

250 mm effective 31 length, influence of the packing immobilization


Ref.

Stationary phase

Mobile phase


Note

Phenols in tobacco smoke (hydroxyquinone, resorcinol, catechol, phenol, isomeric cresols)

Hypersil C 18, 3 ~tm

Acetonitrile-10 mM phosphate buffer pH 9.0 (70:30 or 60:40)

310 mm x 50 gm i.d.


m-Cresol and pyridine

C18 Granocel-14Sh, 7 gm

Acetonitrile-2 mM sodium tetraborate (50:50)

283 mm x 150 gm i.d.


34

2-Phenylmethyl- 1-naphthol

Hypersil C 18, 3 gm

Acetonitrile-20 mM citrate, pH 4-8 (70-80/30-20)

350 mm x 100 gm i.d.


35

2-Phenylmethyl- 1-naphthol


Acetonitrile-50 mM Tris-HC1, 350 mm x 100 gm i.d. pH 8 (80/20)


35

Compound

Phenols

Oxo compounds Acetophenone, propiophenone, butyrophenone, 2',5'-dihydroxyacetophenone, 2',5'-dihydroxypropiophenone

Custom-made macroporous 10 mM Tris + 15 mM boric polyacrylamide-poly(ethylene acid (pH 8.2) with 205 glycol) monolith (acrylate acetonitrile copolymer of acrylic acid, butyl acrylate, methylenebisacrylamide with 3% (w/v) poly(ethylene glycol))

250 mm • 100 ~tm i.d.

205 mm effective length 36

40% Ethyl acrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 4% wt. polymer concentration

Acetonitrile-water (40:60) pH 500 mm x 50 gm i.d. 9.1

Acetone (along with polyaromatic hydrocarbons)

Nucleosil ODS, 5 ~tm

Acetonitrile-0.1 M acetate buffer pH 3.0 (80:20)

250 or 170 mm (nonentrapped or entrapped), i.d. not specified

10 250 and 170 mm are effective column lengths

Acetone, acetophenone, butyrophenone

Linear polymer coated capillary [poly(N-tert.-butyl acrylamide-co-2-acrylamido2-methyl- 1-propanesulfonic acid]

Acetonitrile-50 mM Tris buffer, pH 7.3

750 mm x 25 ~tm i.d.

600 mm effective column length

11

Formaldehyde, acetaldehyde, C18 bonded silica particles, acetone, propionaldehyde, cro- Unimicro 3 ~tm tonaldehyde, methacrolein, 2-butanone, butyraldehyde, benzaldehyde, valeraldehyde, p-tolualdehyde, hexaldehyde

60% Acetonitrile-4% tetrahydrofuran made 5 mM to Tris-HC1 (pH 8.0)or 60% acetonitrile-1 mM borate (pH 9.0) or 60% acetonitrile-4% tetrahydrofuran made 1 mM to borate (pH 9.0)

270 mm x 75 ~tm i.d.

Separated as DNP derivatives, 200 mm packed length

37

Thiourea, benzyl alcohol, Sol-gel bonded 3 ~tm benzaldehyde, benzophenone, ODS/SCX, 80 A pores biphenyl

Acetonitrile-water (80:20) containing 1.5 mMphosphate buffer, pH 3.0

340 mm x 75 ~tm i.d.


28

Alkylphenones

4~

Hydroquinone and its ethers

420 mm to the detector, 6 25~ t,,~~ r~

LiChrospher 100 RP 18, 5 ~tm Acetonitrile (50-70%)-20 mM 300 mm x 100 gm i.d. ammonium acetate


taO


Compound

Stationary phase

Mobile phase


Note

Ref.

320-500 gm i.d.

250-400 mm packed length

36

270 mm x 1O0 gm i.d.

200 mm length

39

200 mm length

39

40

Carbohydrates Malto-oligosaccharides

10 mM Tris + 15 mM boric Custom made macroporous polyacrylamide-poly(ethylene acid (pH 8.2) in acetonitrile0.1% acetic acid (5:95) glycol) matrix

p-Nitrophenyl-c~-D-glucopyra- Zorbax PSM 150, 5 ~tm, nosides and p-nitrophenylconverted to ODS ot-D-malto-oligosaccharides

5 mMNa2HPO4 (pH 6.0)water-acetonitrile (40-45:40-45:20-10)

p-Nitrophenyl-(x-D-glucosides, p-nitrophenyl glucopyranosides, separation of anomers

Zorbax PSM 150, 5 gm, converted to ODS

270 mm x 100 ~tm i.d. 5 mMNa2HPO4 (pH 6.0)water-acetonitrile (42.5-37.5:42.5-37.5:15-25) or 30 mM boric acid (pH 7.0)water-acetonitrile (45-42.5:45-42.5:15-10)

Glucuronides, aromatic

PRP- 1, 10-20 gm

Acetonitrile-2 mM ammonium acetate (pH 7.0) (5:95)

200 mm x 220 jam i.d.

Polyether ketone (PEEK) capillary

Aldopentoses (as 1-phenyl-3methyl-5-pyrazolone derivatives)

Nucleosil silica, in-column 3-aminopropylated or Devosil NH2, 3 ~tm

(25 mM HEPES-NaOH, pH 6.0)-acetonitrile (2" 1)

345 mm x 100 gm i.d.


,~ ~

Monosaccharides of glycopro- DaisoGel silica, in-column teins (as 1-phenyl-3-methylderivatized by 5-pyrazolone derivatives) octadecyltrimethoxysilane or dimethyloctadecyltrimethoxysilylpropylammonium chloride, 3 gm Glycosphingolipids

Porous ODSS, 5 ~tm or non-porous ODSS, 2 ~tm

2.5 mM Phosphate buffer with 345 mm x 100 gm i.d. 5 mM SDS (pH 7.0)acetonitrile (6:4) or 10 mM Tris.HC1 (pH 7.0)-acetonitrile (55:45)


270 mm x 100 gm i.d. Acetonitrile-methanolaqueous sodium borate, pH 9.40 (30:50:20), tetrahydrofuran-aqueous ammonium phosphate, pH 7.00 (80:20), tetrahydrofuran-methanol with 2 mM ammonium acetate (80:20) or tetrahydrofuranaqueous with sodium borate, pH 9.00 (80:20)


~,.~~ t~

r~

Carboxylic acids, simple esters and related compounds Folic acid, p-hydroxybenzoic acid, acetylsalicylic acid, nicotinic acid, thiourea, nicotinamide

Nucleosil 100 3-C 18, 3 gm

Methanol-20 mM disodium tetraborate, pH 8.5 (75:25)

150 mm (effecive length) x 100 gm i.d.

40

Prostaglandins and impurities

Spherisorb ODS-1, 3 ~tm

Acetonitrile-2 mMNa2HPO4, 300 mm x 50 gm i.d. pH 7.3 (75:25)

43

Prostaglandins and impurities

Zorbax SBC8, 3 ~tm

Acetonitrile-10 mM Na2HPO4, pH 9.9 (70:30)

400 mm x 50 gm i.d.

43

ta~

OO


taO OO bO

Compound

Stationary phase

2-Phenylpropionic acid, racemic mixture

Mobile phase


Note

Ref.

trans-3-(3-pyridyl)acrylic acid 50 mMNaH2PO4, pH 4.65 coating molecular imprinting, 3 ~tm

435 mm x 100 ~tm i.d.

350 mm packed length, 44 chiral separation

Phthalate esters

Spherisorb ODS-1, 3 gm

Acetonitrile-50 m M Tris, pH 8.0 (80:20)

350 mm x 100 gm i.d.


45

Malonic acid, sulfate

TSK IC-Anion-SW, 5 gm

Methanol-3 m M phthalic acid, pH 6.8 (adjusted by hexamethylenediamine) (10:90)

289 mm x 50 ~tm i.d.


46

Triglycerides

CEC Hypersil C 18, 3 gm

50 mM Ammonium acetate in 350-500 mm x 100 acetonitrile-isopropanol- ngm i.d. hexane (57:38:5)


47, 48, 49

Alkyl benzoates

40% Ethylacrylate, 50% methacrylic acid, 10% lauryl methacrylate (custom made polymer), 0-3.75 wt. % polymer concentration

Acetonitrile-water (40:60) pH 11.3

500 mm x 50 ~tm i.d.


Phthalate esters

Exsil C18, 1.5 ~tm

Acetonitrile-100 m M Tris, pH 8.6 (90"10)

260 mm x 75 gm i.d.


50

4~

Fatty acids and their phenacyl esters

Hypersil ODS, 3 ~tm

Triglycerides in corn oil

Acetonitrile-50 m M MES [2- 350 or 500 mm x 100 (N-morpholino)ethanesulfonic ~tm i.d. acid], pH 6 (90:10)

250 or 400 mm packed length

51

Hypersil ODS, 3 gm

Acetonitrile-isopropanoln-hexane (57:38:5) 50 m M ammonium acetate

400 mm x 100 gm i.d.

20~

52

Chlorinated alkyl phenoxypropanoates, etc.

Silica gel (Nucleosil, 5 ~tm) coated with Chirasil-Dex

Methanol-20 mM MES buffer, pH 6 (1:1, 2:3 or 7:3)

400 mm x 100 gm i.d.

250 mm effective length, pressuresupported chiral separation

53

Triglycerides in pharmaceutical formulation including testosterone phenyl propionate, testosterone propionate, testosterone isocaproate, testosterone decanoate

Hypersil ODS, 3 gm

Acetonitrile-isopropanoln-hexane (57:38:5) 50 m M ammonium acetate

250 mm x 100 ~tm i.d.

Retinyl esters (C 16, C 17 FA)

Hypersil C18, 3 gm

Lithium acetate in dimethyl formamide (2.5 mM)methanol (99:1)

310-360 mm x 180 gm i.d.


54

Fatty acids and derivatives in food products

CEC Hypersil C18, 3 ~tm

Acetonitrile-50 mM MES, pH 6.0 (90:10)

350-500 mm x 100 ~tm i.d.


48, 49

t~ |

4~

~,,~~

52

TABLE 10.1 (continued) 4~

Compound

Stationary phase

Fatty acid methyl esters (palmitoleic, oleic, eicosenoic and erucic acids)

GROM-SIL ODS-0 AB, 3 ~tm 40 mM Ammonium acetate (pH 9) in water-acetonitrile (50:50)

Thiourea, dimethyl phthalate, diethylphthalate, biphenyl, o-terphenyl


Benzoic acid derivatives RP (Spherisorb ODS-1) and (nitro- and bromo-, positional SAX (Spherisorb SAX) isomers of bromobenzoic acid) packed capillaries, 5 ~tm

Thiourea, methyl p-hydroxybenzoate, ethyl p-hydroxybenzoate, n-propyl p-hydroxybenzoate, n-butyl p-hydroxybenzoate, propranolol, biphenyl, dibucaine

Mobile phase

Acetonitrile-25 mM phosphate, 0.2% hexylamine, pH 2.5 (4:1)


Note

250 mm packed length Coupling with x 100 lam i.d. coordination ion spray mass spectrometry; capillary electrochromatography and pressurized capillary electrochromatography 335 mm x 100 pm i.d.

310 mm x 50 ~tm i.d. 60% Acetonitrile in 2 mM phosphate buffer (pH 2.3) for ODS, 50% acetonitrile in 10 mM phosphate buffer (pH 2.2) for ion-exchange CEC

C18 phases compared (5 lam) Acetonitrile-50 mMTris.HC1, 305 mm x 100 pm i.d. pH 8 (60:40)

Ref. 55



57


r~

Thiourea, dimethyl phthalate, diethyl phthalate, biphenyl, o-terphenyl

ODS-1, 3 pm

Benzoates (methyl- to pentyl-) Grom-Sil ODS-0 ABI, 3 ~tm 4~ ta~

31

Acetonitrile-25 mM Tris.HC1, 250 or 200 mm pH 8 (80:20) effective length x 100 gm i.d.

r,.,~

Acetonitrile-5 mM sodium tetraborate, pH 9.0 (80:20)

370 mm x 22-50 gm i.d.


r~

Steroids

Aldosterone, dexamethasone, 13-estradiol, testosterone

Hypersil C 18, 3 pm or NPS ODS-2, 1.5 pm

Acetonitrile-20 m M sodium acetate, pH 5.0 (70:30) or acetonitrile-25 m M Tris (70:30)

320 mm x 100 gm or 335 mm x 100 gm or 260 mm x 100 pm i.d.

85-250 mm effective length; comparison of frits, double tapers and fritless arrangement

60

Adrenosterone, hydrocortisone, dexamethasone, fluocortolone

Apex ODS, 3 pm

Acetonitrile-5 mM ammonium acetate in water, gradient from 91:9 to 20:80

240 mm x 50 pm i.d.


61

Testosterone, 17-~- and 20-ot-hydroxyprogesterone, androstenedione, progesterone, norethindrone

Hypersil C 18, 3 pm

Acetonitrile-methanol-20 mM Tris-HC1, pH 8.0 (37.5:37.5:25)

350 mm x 100 gm i.d.


62

Tipredane and related compounds


Acetonitrile-50 mM Tris, pH 7.8 (80:20)


Fluticasone and impurities

Hypersil C 18, 3 ~m; Spherisorb ODS-1, 3 ~tm

Acetonitrile-5 mM borate, 400 (or 600) mm x 50 pH 9.0 (80:20) or acetonitrile- pm i.d. 2-10 mMNa2HPO4, pH 8.3-9.9 (75-70:25-30) or acetonitrile-100 mM Tris, pH 9.3 (90:10)

63, 64 43, 65

OO


Compound

Stationary phase

Mobile phase


Note

Ref.

Triamcinolone, hydrocortisone, prednisolone, cortisone, methylprednisolone, betamethasone, dexamethasone, adrenosterone, fluocortolone, triamcinolone acetonide


Acetonitrile-5 mM ammonium acetate, gradient from 17:83 to 38"62

420 mm x 50 pm i.d.


61, 66

Corticosteroids

Hypersil C 18, 3 pm

Acetonitrile-10 to 20 mM phosphoric acid, pH 2.1 (35-95:65-5) or acetonitrile10 mM Tris-HCl, pH 8 (35-95:65-5)

350 mm x 50/100 pm i.d.


67

Corticosterone, testosterone, androsten-3,17-dione, androstan-3,17-dione, pregnan-3,20-dione

Zorbax ODS, 6 pm

Acetonitrile-10 mM borate buffer, pH 8.0 (65:35)

176 m m x 50 p m i.d.

96 mm packed length

68

Aldosterone, hydrocortisone, testosterone



250 mm x 50 ~tm i.d.

Digoxigenin, gitoxigen, cinobufatalin, digitoxigenin, cinobufagin, bufalin


Acetonitrile-4 mM sodium tetraborate, pH 8.0 (70:30) or acetonitrile-water + 0.1% formic acid (70:30)

500 mm x 50 ~tm i.d. or 450 mm x 100 ~tm i.d.

69 250 mm packed length

70 c~

4~ t.,a

Acetonitrile-1.6 mM sodium tetraborate, pH 9.25, 5 mM SDS (60:40)

320 mm x 100 ~tm i.d.

Hydrocortisone, testosterone, 17-~-methyltestosterone, progesterone

Chromspher, nonporous 1.5 ~tm

Estriol, hydrocortisone, estradiol, estrone, testosterone,

Zorbax ODS, 1.8 ~tm

Acetonitrile-0.8 mM sodium 320 mm x 50 jam i.d. tetraborate, 5 mM SDS (80:20)

Spherisorb ODS-1, 3 ~m

Acetonitrile-Tris, pH 7.8 (80:20)

Hydrocortisone, prednisolone, Poly(AMPS-co-IPAAm) hydrocortisone-21-acetate, hydrogel testosterone

100 mM Tris- 150 mM boric acid, pH 8.1


71 ~.,~~

~..~~


72

350 mm • 50 ~tm i.d.

75 mm packed length

73

650 mm x 75 ~tm i.d.


74

Hydrocortisone, prednisolone, Hypersil C18, 3 ~m betamethasone, betamethasone dipropionate, clobetasol butyrate, fluticasone propionate, clobetasone butyrate, betamethasone- 17-valerate

Acetonitrile-2 mM phosphate, 270 mm • 50 ~m i.d. pH 7.8 (80:20)


75

Steroids (neutral or conjugated) and their dansylhydrazine derivatives

Polyacrylic gel-based macroporous particles

Acetonitrile-water-240 mM ammonium formate buffer, pH 3 (55:40:5) (also gradient elution)

350 mm x 100 ~tm i.d.

250 mm active length, laser induced fuorescence detection or coupling with electrospray-ion-trap mass spectrometry

76

Hydrocortisone derivatives



335 mm x 100 ~tm i.d.


26

17-ot-methyltestosterone, 4-pregnen-20c~-ol-3-one, progesterone De-esterified steroids, budesonine, steroid A


ta~

Compound

Stationary phase

Mobile phase


Note

Dexamethasone, betamethasone 17-valerate, fluticasone propionate

Hypersil Duet C 18/SCX mixed mode, 3 gm

Acetonitrile-25 mM ammonium acetate, pH 4.0 (8O:2O)

400 mm x 1O0 pm i.d.

250 mm packed length, 77 coupling with microelectrospray-mass spectrometer

Ref.

Nitro and nitroso compounds

Nitrobenzene, 2,4-dinitroSpherisorb ODS-2, 3 pm toluene and other benzene derivatives along with phenol, phenyl propanol and thiourea N-nitrosodiethanolamine

Acetonitrile-4 mM Tris, pH 435 mm x 50 pm i.d. 9.2 (60:40) or gradient elution to 80:20

C 18 modified etched capillary 0.3 mol/1 Acetic acid containing T-aminobutyric acid (0.375 mol/1), pH 4.41


470 mm x 50 gm i.d.



250 mm packed length, 80 stereoisomer separation

Amines

Fluvoxamine (primary amine) Hypersil ODS, Spherisorb ODS-1, 3 gm

Acetonitrile-phosphate buffer, pH 7.0 (60:40), 6 mM hexylamine

o~

Amino

acids

Dansylated amino acids

Polyacrylamide hydrogel (linear) column, 10 ~tm ethylenechlorotrifluoroethylene

10 m M Tris, 100 m M H3PO4 or water-acetonitrile (10:90) with TFA (0.15%)

Dansylated and non-dansylated amino acids

Dns-L-leucine molecular imprinting

Dansylated phenylalanine

420 mm x 100 gm i.d.


81

Acetonitrile-acetic acid-water 500 mm x 50 gm i.d. (90:5:5)

250 mm packed length, chiral separation

82

Dns-L-phenylalanine molecular imprinting

Acetonitrile-10 m M phosphate, pH 7 (10:1)

1 0 0 0 m m x 25 ~ m i.d.


83

Dansylated phenylalanine

Dns-L-phenylalanine molecular imprinting

Acetonitrile-100 m M acetate (80:20)



10

DNP amino acids

[3-CD bonded silica particles, 5 pm

Methanol-10-20 m M triethylamine acetate buffer, pH 4.71 (10-25:90-80)

400 mm x 50 pm i.d.


84

PTH amino acids

Zorbax ODS, 3.5 pm

Acetonitrile-5 m M phosphate, 207 mm x 50 pm i.d. pH 7.55 (gradient from 30:70 to 60:40)


68

PTH amino acids

Monolithic packing of sintered Zorbax ODS, 6 pm

Acetonitrile-5 m M sodium phosphate, pH 7.5 (30:70)

330 mm x 75 pm i.d.

The sorbent was reoctadecylated; 230 mm packed length

85

PTH amino acids

Zorbax ODS, 3.5 gm

Gradient elution (A: 2 m M aqueous ammonium acetate, pH 7.0; B: 2 m M a m m o n i u m acetate in water-acetonitrile 1:9) from 30 to 90% B

1 5 0 m m x 75 p m i.d.

coupling with electrospray ionization and time-of-flight mass spectrometry

86

4~

t~

ta,9


Stationary phase

Mobile phase

Phenylalanine, tyrosine, phenylglycine, tryptophan, serine

L-Phenylalanine anilide molecular imprinting

Acetonitrile-acetic acid-water 500 mm x 75 gm i.d. (80:10:10)


Phenylalanine, tyrosine, p-fluorophenylalanine, phenylglycine, phenylalanine amide, DOPA

Phenylalanine or phenylalanine amide molecular imprinting

Acetonitrile-acetic acid-water 400 mm x 75 pm i.d. (90:5"5)


N-derivatized amino acids (DNZ, FMOC), their enantiomers

Silica gel (Kromasil) modified with a basic tert.-butyl carbamoyl quinine chiral selector, 3 ~tm

Acetonitrile or methanol-50 mM acetic acid (80:20), pH 6 (titrated with triethylamine or ammonia)

335 mm x 100 (or 75) pm i.d.

Acetonitrile-5 mMphosphate buffer, 7 (80:20)

300 mm (active length) Enantiomer separation x 100 pm i.d.

N-(3,5-dinitrobenzoyl)leucine Monoliths (2-hydroxyethyl diallyl amide methacrylate carbamate copolymerized with four different chiral selectors)


Note

Ref.

Compound

250 mm packed length, 89 chiral separation by weak anion-exchange type chiral stationary phase 90

N-derivatized amino acids, their enantiomers

CEC-Hypersil, 3 ~tm

r

| 4a.

Non-aqueous: methanol, 20 mM ammonium acetate, 80 mM acetic acid, 5 mM tert.butyl carbamoyl quinine; Aqueous: methanol-100 mM ammonium acetate (80:20), pH 6.0, 5 mM tert.-butyl carbamoyl quinine

335 mm x 100 gm i.d.

250 mm effective 91 length, chiral separation

,.,,, ~ t% ~,,~~

Phenylthiohydantoin-amino acids

Chromspher ODS, 1.5 gm

Phosphate buffer pH 7.2, 5 mM SDS, 5% acetonitrile and 5% tetrahydrofuran

325 mm x 100 lam i.d.


Dansylated amino acids

]3-Cyclodextrin-bonded positively charged polyacrylamide gel

200 mM Tris, 300 mMboric acid buffer, pH 8.1

550-700 mm x 75 jam i.d.


Benzyloxycarbonyl, N-(3,5dinitrobenzyloxycarbonyl), 9-fluorenylmethoxycarbonyl, benzoyl, acetyl and N-(2,4dinitrophenyl) derivatized amino acids and profens

WAX (weak anion-exchange) Acetonitrile-methanol (80"20)+400 mM acetic type CSP (tert.acid+4 mM triethylamine butylcarbamoylquinine as chiral selector on Hypersil silica gel), 3 gm

335 mm x 100 ~tm i.d.


Tryptophan and dinitrobenzoyl leucine enantiomers

Chirobiotic T (teicoplanin), 5 gm

Acetonitrile-2 mM (or 5 mM) 330 mm x 100 gm i.d. Na2HPO4, pH 7, 3.5 or 2.3


DNZ-leucine, DNB-leucine, Fmoc-Leucine

Silica based (Kromasil 1005gm, Hypersil 120-3gm, Micra NPS-1.5~tm) weak anion-exchange-type chiral stationary phases

Acetonitrile or methanol-100 mM MES (80:20)

335 mm x 75 or 100 gm i.d.

92



Compound

to

Stationary phase

Mobile phase


Note

Ref.

60 mm packed length

97

Peptides, proteins

Peptide map, cytochrome c tryptic digest

ODS, 1.5 ~m, 100 A pore size A: 0.1% TFA in water, pH 2, (Polymicro Technologies) B" as A containing 50% (by vol.) acetonitrile; gradient 0-100% B over 10 min

200 mm x 100 ~tm i.d.

Nonapeptide

Nucleosil 100-5C8, 3 gm

Methanol-4 mM ammonium acetate, pH 8.0 (40:60)

250 mm x 220 ~tm i.d.

98

Cytochrome c, bovine, tryptic Vydac C 18, 3 ~tm digest, chicken albumin, tryptic digest

Acetonitrile-0.07% TFA in water (gradient from 0:100 to 40:60)

60 mm x 180 ~tm i.d.

99

Vydac C 18, 3 ~tm Bradykinin, angiotensin II, angiotensin I, Met-enkephalinArg-Phe, neurotensin

0.07% TFA in wateracetonitrile (75:25)

120 mm x 180 ~tm i.d.

99

100

[~-Lactoglobulin, bovine

Vydac C18, 3 ~tm

0.04% TFA in acetonitrilewater (gradient from 0:100 to 75:25)

60 mm x 180 ~tm i.d.

Tetrapeptides, pentapeptides

Gigaporous PLSCX, 8 gm

Acetonitrile-25 mM phosphate, pH 3.5 (40:60)

3 7 0 m m x 180~tmi.d.


101

Tyrosine-containing peptides

Polyacrylamide poly(ethylene Acetonitrile-10 mM Tris, glycol) macroporous packing 15 mMboric acid, pH 8.2 (47:53)

250 mm x 100 gm i.d.


36

Lysozyme, angiotensins r r~

C 18, etched and modified column; diol open tubular column

30 m M phosphate, pH 2.14, 30 m M citric acid, pH 3-25 m M [3-alanine, 30 mM acetic acid, pH 4.14

450 mm x 50 pm i.d.


102 ~,~~ t~

4~ too

Cytochrome c, lysozyme, myoglobin, ribonuclease A

C 18, etched and modified column; diol open tubular column

30 m M phosphate, pH 2.14, 30 m M citric acid, pH 3-25 m M [3-alanine, 30 mM acetic acid, pH 4.14

450 mm x 50 pm i.d.


103

Enkephalin methylester, enkephalin amide

Gromsil ODS-2, 1.5 pm

Acetonitrile-0.07 ml/1 TFA in water (80:20)

400 mm x 100 pm i.d.


104

Tetrapeptide, C- and N-protected


Acetonitrile-50 mM Tris, pH 7.8 (80:20)

335 mm x 50 gm i.d.


63, 64

Mixture of peptides and proteins (lysozyme, angiotensin I and III, bradykinin, ribonuclease A)

C 18-modified etched capillary 30 m M citric acid-24.5 m M [3-alanine, pH 3.0



Lysozymes

Diol or C 18-modified etched capillaries

30 mMPhosphate, pH 2.14 or 450 mm x 50 ~tm i.d. 30 m M citric acid-19 m M Tris, pH 3.0 for C 18 capillary or 30 m M acetic acid-25 m M [3-alanine, pH 4.41 for diol capillary


Angiotensins

Diol or C 18-modified etched capillaries

30 mMphosphate, pH 2.14


450 mm x 50 pm i.d.


Mobile phase


Compound

Stationary phase

Carbonic anhydrase, myoglobin, amylase, trypsinogen, ribonuclease A, chymotrypsinogen A, mesityl oxide

Polyaspartic acid immobilized Salt or pH gradient elution in 400 mm x 75 pm i.d. phosphate buffer (pH 6.0) or on the capillary wall isocratic elution with 100 mM NaC1 in 10 mM phosphate (pH 6-8)

Peptide mapping ovalbumin, tryptic digest

C 18 COMOSS microfabricated column

Acetonitrile-10 mM potassium phosphate, pH 9.0 (1:3)

Oxytocin, desmopressin, cerbetocin and related synthetic peptides

CEC Hypersil C8, CEC Hypersil C 18, 5 gm, Spherisorb mixed mode C 18/SCX phase 3 pm

Triethylamine buffers, pH 3.0 335 mm • 100 pm i.d. adjusted with phosphoric acid with different proportion of acetonitrile

Basic proteins and peptides

0, 10, 20, 30% Acetonitrile in 470 mm x 20 pm i.d. PLOT (porous-layer open 20 mM aqueous sodium tubular) column, i.e., fused ilica capillary with 2 gm thick phosphate, pH 2.5 polymer layer (highly crosslinked in situ polymerized vinylbenzyl chloride and divinylbenzene)

1.5 pm wide, 10 ~tm deep rectangular channel, 45 mm long

Note

Ref.


107

700 s needed for run completion

108


109


Recombinant human growth hormone, tryptic digests o~

Vydac 218TPB5 (C18), 5 gm, A: 0.1% TFA/water, B: 300 A 0.09% TFA/acetonitrile, gradient from 0 to 60% B

250 mm x 100 gm i.d.

4~

Electrically assisted 111 capillary HPLC, coupling with electrospray ionizationmass spectrometry

Cytochrome c

Capillary etched with liquid crystals (cholesteryl or cyanopentoxy modified)

60 m M Citric acid and 50 mM 700 or 500 mm x 50 13-alanine, pH 3.00 gm i.d

450 or 250 mm effective length, open tubular CEC

112

Lysozymes, cytochromes c, aspartame

Silica capillary was etched and chemically (C18) modified

pH 3.7

515 or 580 mm x 20 l.tm i.d.

220 or 260 mm effective length, open tubular electrochromatography

113

Ribonuclease A, insulin, c~-lactalbumin, myoglobin

Methacrylic monolith with tertiary amino functions

30% Acetonitrile in 60 m M aqueous sodium phosphate, pH 2.5



Hypersil ODS, 3 ~tm

30% Methanol, 10% acetonitrile in 5 m M aqueous ammonium acetate

250 mm • 25 gm i.d.

CEC/MS (quadrupole) coupling

Acetonitrile-5 m M acetic acid, 2-3 mM triethylamine, pH 5.0 buffer (8:92)

250 mm • 100 ~tm i.d.

effective column length 116 not specified, 20-26~

Nucleotides, nucleosides Adenosine-styrene oxide adducts, neutral and positively charged; inosine-styrene oxide adducts, neutral and positively charged

CEC Hypersil C 18, 3 gm Nucleosides (adenosine, cytidine, guanosine,thymidine and uridine)

115

-m,

taO


O',

Compound

Stationary phase

Mobile phase


Note

Ref.

Purine and pyrimidine bases and their nucleosides (adenosine, adenine, cytidine, cytosine, guanosine, guanine, inosine, thymine, uridine, uracil)

ODSS (octadecyl sulfonated silica), 10 gm

Acetonitrile-4.8 m M sodium acetate, pH 4.5 (40:60)

270 mm x 20.5 gm i.d


AMP, ADP and ATP

Nucleosil 100-C 18, 5 gm

Methanol-2 m M dibutylamine, pH 5.0 (10:90)

200 mm x 220 gm i.d.

118

d(GATGCATAGG-OH) and by-products, dC3-dC 11

Gromsil ODS-2, 5 gm

Acetonitrile-10 m M triethylamine acetate (12:88), acetonitrile-10 m M ammonium acetate, gradient from 0"100 to 10:90

500 mm x 50-100 gm i.d.


119

Thymine, cytosine, adenine, guanine, adenosine

Capillary etched with liquid crystals (cholesteryl or cyanopentoxy modified)

60 mM Phosphoric acid and 38 m M Tris, pH 2.14

700 or 500 mm x 50 ~tm i.d

450 or 250 mm effective length, open tubular CEC

112

Nucleosides (adenosine, cytidine, guanosine, uridine, thymidine)

Hypersil Phenyl, 3 ~tm, 120 A Acetic acid-ammonia, pH 5acetonitrile (95:5)

270 mm x 75 ~tm i.d

200 or 70 mm bed length

120

Alkaloids

Morphine alkaloids 4~

~.,~~

Nucleosil 100-C 18, 5 ~tm

Acetonitrile-2 m M ammonium acetate (40:60)

150 mm x 220 gm i.d.

118

YMC C30, 5 gm

N,N-Dimethylformamideacetonitrile-methanol (29:70"1) with 2.5 m M lithium acetate

360 mm x 180 ~tm i.d.


Macrocyclic antibiotics (lactone, from Streptomyces $541)

CEC Hypersil, 3 gm

Acetonitrile-5 m M borate, pH 9 (80:20)

430 mm x 50 gm i.d.

122

Tetracyclines

Etched and modified C 18 open tubular column

Methanol-30 m M citrate containing 24.5 m M 13alanine, pH 3.0 (40:60)

250 mm x 50 gm i.d.

123, 124

Cephalosporins (cefuroxime, axetil)

CEC Hypersil, 3 lam

Acetonitrile-5 m M borate, pH 9 (80:20)


Cephalosporins (cefuroxime, axetil)


Acetonitrile-10 m M Na2HPO4, pH 9.5 (80:20)


Tetracyclines

C 18-modified etched capillary 30 m M phosphate-19 mM Tris, pH 2.14, 40% methanol

Vitamins

|

4~

Retinyl esters

Antibiotics

450 mm x 50 gm i.d.

For MS assay

43 43


--..I


Stationary phase

Mobile phase


Note

Ref.

Triazine herbicides

Hypersil C 18, 3 pm, Hypersil C8, 3 gm, Spherisorb C6/SCX, 3 ~tm

Acetonitrile-25 m M sodium acetate, pH 8 (50:50)

335 mm x 100 pm i.d.


30

Primicarb


Acetonitrile-20 m M aqueous Tris, pH 9.0 (60:40)

330 mm x 50 pm i.d.


50

Pesticides

Hypersil ODS C 18, 3 ~tm

Acetonitrile-25 m M H E P E S buffer, pH 7.55 (75:25)

490 mm x 100 pm i.d.


Urea herbicides

Zorbax ODS, 5 ~tm

270 mm x 100 pm i.d. Methanol-5 mMNaH2PO4, pH 6.0 (70:30) or acetonitrile5 mMNaH2PO4, pH 6.0

200 mm effective 126 length; preconcentration

Compound Pesticides

(5o:5o) 245 mm packed length

127

335 mm • 100 ~tm i.d. Acetonitrile-20 m M sodium dihydrogen phosphate, pH 4.0 (60:40)


128

Acetonitrile-13 m M T F A , pH 3.5 (60:40)


60

Primicarb

CEC Hypersil C18, 3 ~tm

Acetonitrile-5 m M aqueous Tris, pH 8.6 (42:58, 12:88)

Cinosulfuron

Synchropak (C 18), non-endpacked

Cinosulforon and by-products

Gromsil ODS-O AB, 3 gm

330 mm x 50 pm i.d.

337 mm x 75 pm i.d.

c~

Polychlorinated dibenzo-pdioxins

4~ i

4~

ODS C 18, 3 pm

Spherisorb ODS-1, 3 gm, Herbicides (desisopropylaimmobilized trazine, desethylatrazine, simazine, cyanazine, atrazine, sebutylazine, propazine, terbutylazine, 2-hydroxyterbutylazine, 2-hydroxyatrazine)

Acetonitrile-25 mM Tris, pU 8.5 (75-80"25-20)

450 mm x 100 ~tm i.d.

Acetonitrile-25 mM Tris.HCl buffer, pH 8 (56:44)

335 mm x 100 pm i.d.

250 mm effective 31 length, influence of the packing immobilization

Channel depth 5.2 lam

length not specified, chip arrangement

130

Polyether ketone (PEEK) capillary

40


129 r..~~

Dyes Coumarin dyes (C440, C450, C460, C480)

Octadecylsilane or 10 mM Borate with 29% octadecyldiisobutyl(dimethyl- acetonitrile, pH 8.4 amino)silane surface modified capillary

Food dyes (E 102, E 110, E122, E123, E124)

Nucleosil 10-5C 18, 5 gm

Textile dyes (azo- and anthra- Hypersil C 18, 3 gm quinone compounds) Carotenoid isomers

Methanol-10 mM ammonium 200 mm x 250 pm i.d. acetate, pH 8.5 (20:80) Acetonitrile-4 mM borate, pH 8.0 (80:20)

Polymeric C30 (Rainin 30 nm Isocratic: methanol-methylpore size or ProntoSIL 30 nm tert.-butyl ether-buffer pore size), 3 pm (35:60:5), acetone-buffer (99:1, 95:5, 90:10, 85:15) when buffer is 1 mMborate buffer. Gradient elution: acetone-1 mM sodium borate buffer, gradient from 80:10 to 99:1

25 mm x 75 pm i.d. 250 mm x 98 gm i.d.

131 330 mm total length

132

~..,~

4~ O


Compound

Drugs (for steroid drugs s e e

Stationary phase also

Ref.

Mobile phase


Note

Acetonitrile-buffer, pH 2.5 (75"25)

340 mm x 100 lam i.d.

250 mm to the detector, 133 temp. 25~ comparison with HPLC and CZE

Steroids)

Drugs of abuse: amphetamine, CEC Hypersil C8, 3 pm metaphetamine, procaine, cocaine, heroin, quinine, noscapine and thiourea Drugs of abuse: phenobarbital, testosterone, cannabinol, testosterone propionate, A9-tetracannabinol, A9-tetracannabinolic acid

CEC Hypersil C8, 3 pm

Acetonitrile-25 mM phosphate buffer, pH 2.5, gradient from 60:40 to 75:25

340 mm x 100 pm i.d.

250 mm to the detector, 133 comparison with HPLC and CZE

Dexamethasone, hydrocortisone, prednisolone acetate, cortisone acetate

Hypersil C8, 3 lam


370 mm x 50 pm i.d.


Hydrocortisone, hydrocortisone- 17-valerate, hydrocortisone- 17-butyrate, betamethasone- 17-valerate, clobetasol- 17-propionate and preservatives (parabenes, benzyl alcohol, sorbic acid, chlorocresol)

Innovatech SCX/C 18, Hypersil ODS-1, ODS-2, 3 pm

250 mm x 50 ~tm i.d. Acetonitrile-50 mM sodium dihydrogen phosphate, pH 3.5 (30:70)

134

135

"x

4~

Acidic and basic drugs analysed simultaneously (amphetamine, metamphetamine, procaine, cocaine, heroin, quinine, noscapine)

CEC Hypersil C8

Acetonitrile-50 mM phosphate buffer, pH 2.5 (75:25) (75, 60, 45 and 30% acetonitrile also tested)

340 mm x 100 ~tm i.d.


~,~~ r~

ta, a |

4~

~,~~

Acidic and basic drugs CEC Hypersil C8 (phenobarbital, diazepam, methaqualone, testosterone, cannabinol, testosterone propionate, 9-tetrahydrocannabinol, 9-tetrahydrocannabinolic acid)

Acetonitrile-50 mM 340 mm x 100 ~m i.d. phosphate buffer, pH 2.5 (60:40) with 2 gl/ml hexylamine for 1 min, then linear gradient to acetonitrile25 mM phosphate buffer with 2 gl/ml hexylamine (75:25)

250 mm to the detector, 136 stepped gradient, comparison with MECC

Acidic and basic drugs (see above)

CEC Hypersil C8

Acetonitrile-phosphate 340 mm x 100 gm i.d. buffer, pH 2.0 with 3.4 gl/ml hexylamine (2:98) for 20 min, then linear gradient to acetonitrile-phosphate buffer with 3.4 lal/ml hexylamine (65:35) and hold for 10 min

250 mm to the detector, 136 complex stepped gradient

Tripredane and related compounds

Spherisorb ODS-1, 3 lam


Barbital, butethal, phenobarbi- Hypersil C8, Hypersil C 18, tal, amylobarbital, secoHypersil phenyl, 3 tam barbital, hexobarbital

250 mm x 50 gm i.d.

(a) Acetonitrile-50 mM MES, 350 mm x 100 ~tm i.d. pH 6.1-water (60-40:20:2040); (b) methanol or acetonitrile50 mM phosphate, pH 4.5water (40-50:20:40-30)


137, 138


O b,.)

Mobile phase


Stationary phase

Bendroflumethiazide

Spherisorb ODS-1, Spherisorb Acetonitrile-10-50 mM 500-550 mm • 50 ~tm 260-350 mm to the detector SCX, 3 ~tm NaH2PO4, pH 3.5-9.8 (70:30) i.d.

Barbiturates

Permethyl-[3-cyclodextrinMethanol-5 mM phosphate modified silica gel (Nucleosil, buffer, pH 7.0 (1:4) 5 ~tm)

Chlorthalidone, hydroflumethiazide, bendroflumethiazide, bumetanide

Spherisorb ODS- 1

Acetonitrile-5 mMNaH2PO4, 330 mm x 100 pm i.d. pH 2.3 (gradient from 40:60 to 60:40)

Hydroflumethiazide, methylchlothiazide, metolazone, epitizide, bendrofluazide


Acetonitrile-5 mM ammonium acetate in water, gradient from 50:50 to 20:80

Bendroflumethiazide, nortriptyline, chlomipramine, methdilazine, imipramine, desipramine


550 mm x 50 ~tm i.d. Acetonitrile-10-50 mM Na2HPO4, pH 5.7-9.8 (70:30)


65

Bendroflumethiazide, nortriptyline, chlomipramine, methdilazine, imipramine, desipramine

Spherisorb SCX, 3 gm

Acetonitrile-50 mM Na2HPO4, pH 3.5 (70:30)

500 mm x 50 ~tm i.d.


65

400 mm x 100 gm i.d.

Note

Ref.

Compound

65

235 mm effective length, chiral separation, pressuresupported CEC

139


140

61, 66

460 mm x 50 ~tm i.d.

Nitrazepam, diazepam

4~

Cloxazolam, nitrazepam, clotiazepam, diazepam

460 mm x 50 gm i.d.

Hypersil C 18, 3 jam

5 mM Ammonium acetate in acetonitrile-water, gradient from 50:50 to 80:20

Etched cholesterol modified open tubular column

Acetonitrile-10 mMTris-HC1, 700 mm x 50-75 gm pH 7.3 (0-20:100-80) i.d.

66 r..~,


141

r~

|

4~

Nonsteroidal antinflammatory CEC Hypersil C18/SCX, 3 ~tm Acetonitrile-50 mM drugs (ketoprofen, naproxen, Na2HPO4-water (60:20:20) flurbiprofen, indomethacin, ibuprofen) Bumetanide, flurbiprofen, p-hydroxybenzoic acid

Hypersil C 18 or Hypersil C 18/SCX, 3 gm

Acetaminophen, caffeine, ace- GROM-SIL1 100 ODS-0 AB, tylsalicylic acid 5 gm (Thomapyrin |

Antiviral drugs (suramin)

Nucleosil 100 C 18, 5 ~tm

Bare silica Micra, 3 ~tm Amino group containing drugs (codeine phosphate, ephedrine hydrochloride, thebaine, berberine hydrochloride, jatrorrizine hydrochloride, cocaine hydrochloride)

Acetonitrile-50 mM Na2HPO4, pH 2.3-2.5-water (40:20:40)

210 mm x 50 ~tm i.d.

138

210-230 mm x 50 ~tm i.d.

63, 138

200 mm x 250 ~tm i.d. 2 mM Borate in D 2 0 deuterated acetonitrile (80:20) or 1 mMborate in D 2 0 deuterated acetonitrile (gradient from 100:0 to 70:30) Methanol-2 mM dibutylamine, pH 5.0 (10:90)

200 mm x 220 gm i.d.

Acetonitrile-5-20 mM TrisHC1, pH 7.5-10 (90:35-10)

270 mm x 65-75 ~tm i.d.

coupling with NMR

142


143

4~ O

4~


Compound

Stationary phase

Mobile phase


Isradepin and and byproducts

Hypersil C18, 3 ktm


143 mm x 50 ktm i.d.

144

Methanol-5 mM borate buffer, pH 8.5 (60:40)

260 mm x 50 ~m i.d.

145

Methanol-4 mM ammonium acetate, pH 5 (40:60)

235 mm x 220 ~tm i.d.

98

NPS ODS-2, 1.5 ~tm Cardiac glycosides (digoxigenin, digoxin, digitoxigenin)

Acetonitrile-25 mM Tris, pH 8 (50:50) made 0.3% to SDS

260 mm x 100 ~tm i.d.

Nucleosil 5C8, 3 gm 2-Phenylethylamine derivatives (epinephrine, DOPA, 2-amino-3-hydroxy-3-phenylpropanol, ephedrine)

Methanol-5 mMNaH2PO4, pH 3.1 (60:40)

155 mm • 100 jam i.d.

Antiepileptic drugs (ethosuc- Spherisorb ODS-1, 3 ~tm cinimide, primidon, carbamazepine- 10,11-diol, carbamazepine- 10,11-epoxide, carbamazepine, phenytoin Sulfanilamide, sulfaflurazol, sulfadicramide

Nucleosil 100-5C8, 3 gm

Note


Ref.

60

145

c~

Chiral drugs and chiral compounds (clomipramine, ~ chlorodiazepoxide, diazepam, .~ temazepam, doxepine, terbu"~" taline, clenbuterol, homota~ k atropine, tropicamide, ,~ 3,5-dinitrobenzoyl alanine methyl ester) o~

450 mm x 50 gm i.d. Silanized/hydrosilylated inner Phosphate pH 2.14, Tris surface of the capillary pH 3.0, citric acid + 13-alanine pH 3.7, lactic acid + 13-alanine pH 4.41, ,/-aminobutyric acid pH 6.0 (38-60 mM)

PM-13-CD, 5 gm Barbiturates (mephobarbital, hexobarbital, pentobarbital, 1-methyl-5-(2-propyl)-5(n-propyl)barbituric acid, 5-ethyl- 1-methyl-5-(n-propyl) barbituric acid), benzoin, c~methyl-ot-phenylsuccinimide, gluthethimide, methylthiohydantoin-proline, methyl mandelate

Methanol-5 mM phosphate, pH 7 (20:80)

400 mm x 100 9m i.d.

250 mm effective length, open tubular chromatography, chiral separations

146 .,,,o

~o r~


147


Hexobarbital

13-CD, 5 pm

Acetonitrile-4 mM phosphate, 400 mm x 50 ~tm i.d. pH 6.8 (5:95), methanol-5 mM triethylamine acetate, pH 4.71 (15:85)

Chlorthalidone

Spherisorb $30DS-1, 3 gm

10 mM HP-I3-CD in acetonitrile-5 mM phosphate, pH 6.5 (20:80)

520 mm x 50 ~tm i.d.


148

Chlorthalidone

Hydroxypropyl-[3-CD bonded Acetonitrile-5 mMphosphate, 580 mm x 50 gm i.d. silica, 5 ~tm pH 6.5 (25:75)


148


Compound

Stationary phase

Mianserin

Hydroxypropyl-13-CD bonded Acetonitrile-10 mM silica, 5 pm phosphate, pH 7.5 (50:50)

Ibuprofen, cicloprofen, 1-phenylethanol, 1,1'binaphthyl-2,2'-dihydrogenphosphate, flurbiprofen, carprofen, etodolac, warfarin, hexobarbital

Chirasil-Dex coated column, 0.2 lam

Alkylated barbituric acids, 6chloro-c~-methylcarbazole-2acetic acid, 1-phenylethanol, 1-phenyl- 1-propanol, otmethy 1-2,3,4,5,6-pentafluo robenzyl alcohol, 1-(2naphthyl)-ethanol,

Mobile phase


Note

Ref.



148

20 mM Borate-phosphate, pH 800 mm x 50 pm i.d. 7, or 20 mM Tris-HC1, pH 7

Chiral separation

149, 147, 150, 151

Chirasil-Dex coated column, 0.15 pm

Acetonotrile-20 mM borate700-980 mm x 50 ~tm phosphate buffer, pH 7 i.d., or 620 mm • (10:90) or methanol-20 mM 25 gm i.d. borate-phosphate buffer, pH 7 (38-3:62-97)

600-880 mm packed 152 length, chiral separation or 500 mm packed length, chiral separation

ODS C18, 5 gm

20 mM Sodium phosphate, pH 3.0-12 mM 13-CD-4-5 mM sodium 1-heptanesulfonate

170-230 mm packed 153 length, chiral separation

1-(p-biphenyl)-ethanol Salsolinol

200-290 mm x 75 gm i.d.

c~

Hexobarbital, pentobarbital, isofamide, cyclophophamide, disopyramide

c~1-Acid glycoprotein-bonded 1-Propanol-5 mM phosphate, silica, 5 ~tm pH 6.5 (2:98) or 2-propanol2 mM phosphate, pH 5.5 (2:98) or 2-propanol-4 mM phosphate, pH 6.5 (2:98)

420 mm x 50 gm i.d.


154

Temazepam, benzoin

Human serum albuminbonded silica, 7 gm

2-Propanol-4 mM phosphate, pH 7 (5-7.5:95-92.5)



155

Hexobarbital, warfarin

Vancomycin-coated silica, 3 lam

Acetonitrile-0.1% triethylamine acetate, pH 4 (80:20) or acetonitrile-0.1% triethylamine acetate, pH 5 (2O:8O)

500 mm x 1O0 gm i.d.


156

Glutethimide, 1-(9-anthryl)2,2,2-trifluoro-ethanol, mephobarbital, aminoglutethimide

3,5-dimethylphenylcarbamoyl Acetonitrile-40 mM cellulose and p-methylphosphate, pH 7 (15-40:85benzoyl cellulose coated 6O) column, 0.25 gm

570 mm x 50 gm i.d.


157

Acebutolol, alprenolol, atenolol, metoprolol, pindolol, prenalterol, propranolol

(R)-propranolol molecular imprinting phase

Acetonitrile-100 mM phosphoric acid and triethanolamine to pH 3.0 (8O:2O)

350-900 mm x 75 gm i.d.

265-815 mm packed 158 length, chiral separation

Propranolol, metoprolol

(R)-propranolol or (S)metoprolol molecular imprinting phase

Acetonitrile-2-4 mM acetate, pH 3.0 (80:20)



4~ t~ !

4~

159

4~


Compound

Stationary phase

Mobile phase


Note

Ref.

Terbutaline, benzoin

Charged polyacrylamide gel

50 mg/ml Poly [3-cyclodextrin, 50 mg/ml carboxymethyl ~-cyclodextrin, 5% Tween 20, 200 mM Tris, 300 mMboric acid buffer, pH 9.0

700 mm x 75 pm i.d.


Terbutaline, propranolol, benzoin

[3-Cyclodextrin-bonded charged polyacrylamide gel

200 mM Tris, 300 mM boric acid buffer, pH 9.0

700 mm x 75 pm i.d.

161 350 mm effective length, chiral separation

Benzodiazepines

Electropak phenyl, 3 pm

Tris.HC1, pH 8-acetonitrile (40:60)

470 mm x 75 pm i.d.


Benzoin acetate, methylbenzoin, Tr6ger's base, trans-stilbene oxide and 1, l'-binaphthyl-2,2'-diol

Aminopropyl silica gel (LiChrospher 1000, 5 gm) coated with helically chiral poly(diphenyl-2pyridylmethyl methacrylate) (PDPM)

Methanolic solution of ammonium acetate (2.5 mM, pH 4.5)


163 200 mm effective length, chiral separation

Mephobarbital, hexobarbital, MTH proline, methyls of mecoprop, diclofop and fenoxaprop, barbiturates, chlorinated alkyl phenoxypropanoates, etc.

Silica gel (Nucleosil, 5 gin) coated with Chirasil-Dex

Methanol-20 mM MES buffer, pH 6 (1"1, 2:3 or 7:3)

290 (400) mm x 1O0 lam i.d.

200 (250) mm effective 53 length, pressuresupported chiral separation

162

o~

"x I%

4~

Opiate drugs

C 18, 1.5 ~tm

Thalidomide and its metabolites

LiChrospher 100 RP- 18, 5 pm 5 mM Acetonitrileammomium acetate, pH 6 (60:40)

too

10mMTris, 5mMSDS, 20% 280 mm x 75 tam i.d. acetonitrile, pH 8.3 330 mm x 100 lam i.d.

150mmeffectivelength 164 245 mm packed length, chiral separation

165 r~

Neutral polar pharmaceuticals Reversed-phase non-porous silica (NPS C-18, NPS ODS1, NPS PolyEncap B 1, NPS TAS- 1), 1.5 ~tm

Acetonitrile-2 mMphosphate, 335 mm x 75 pm i.d. pH 3.0 (60:40, 50:50 or 40:60)

250 mm effective length; comparison of various 1.5 lam nonporous phases

166

Benzodiazepines (temazepam, Capillary etched with liquid oxazepam, clonazepam, diaze- crystals (cholesteryl) pam, nitrazepam)

60 mM Phosphoric acid and 38 mMTris, pH 2.14

700 mm x 50 pm i.d

450 mm effective length, open tubular CEC

112

Theophylline, caffeine, acetaminophen, hydrochlorothiazide, 13-hydroxyethyltheophylline, phenylbutazone and theobromine

Silica EP-75-20-3-Si, 3 ~tm

Isopropanol-hexane-1 mM Tris (52:40:8, pH 8)

270 mm x 75 pm i.d.


Thalidomide and its metabolites

Aminopropyl silica coated Methanol-ethanol (75:25) with Chiralpak AD and/or OD containing 2.5 mM ammomium acetate

335 mm x 100 gm i.d.


Clenbuterol, salbutamol, methadone, polynuclear aromatic hydrocarbons

MOS Hypersil, 3 gm

25 mM Ammonium acetate 160 m m x 75 p m i.d. buffer, pH 5.0, in acetonitrilewater (90:10) or 2 mM Tris, pH 8.0 in acetonitrile-water (80:20)

169


Compound

Stationary phase

Mobile phase


Note

Ref.

Indapamide, lormetazepam, transstilbene oxide, benzoin

3,5-Dimethylphenylcarbamoyl cellulose immobilized on silica gel, 7 pm

6.6 or 0.8 mM citrate buffer (pH 5 or 6)-acetonitrile (30:70, 40:60 or 50:50)

270 mm x 100 lam i.d.


[3-adrenergic blocking agent and various enantiomers

Vancomycin bonded on LiChrospher diol silica, 5 pm

Methanol-acetonitrile-acetic 355 mm x 75 l.tm i.d. acid-triethylamine (80-20:2080:0.1-0.3:0.1-0.4)


Tricyclic antidepressants

Custom made molecular imprint polymer sorbents

330 mm x 100 ~tm i.d. Acetonitrile-10 mM sodium acetate, pH 3.0 (98:2) with 0.02% trifluoroacetic acid and 0.015% triethylamine


Model drug mixture (procaine, timolol, ambroxol, metoclopramide, thiourea, naproxene, antipyrine)

Spherisorb ODS-1, 3 lam, CEC Hypersil C 18, 3 pm or CEC Hypersil C 8 , 3 lam

Acetonitrile-25 mM phosphate, 0.2% hexylamine, pH 2.5 (80:20)

335 mm x 100 pm i.d.


Carbovir, ranitidine, ondansetron, imipramine, amitriptyline, clomipramine

Water symmetry shield RP-8, 3.5 ~tm


330 mm x 50 pm i.d.


Anti-inflammatory drugs, non-steroidal (etodolac and derivatives)

330 mm x 100 gm i.d. LiChrospher 100 RP-18, 5 gm Acetonitrile-75 (10) mM ammonium formate, pH 2.5 or 3.0 (40:60 or 50:50)

172

173

245 mm packed length, 174 coupling with electrospray ionisationmass spectrometry

~--

Drug test mixture (aminoChiralcel OD, glutethimide, 2,2'-diaminoChiralpak AD, 6,6'-dime thylbiph en yl econa- Chiralcel OJ zole, etozolin, glutethimide, indapamide, metomidate, piprozolin, trans-stilbene oxide, Tr6ger's base)

Methanolic or ethanolic ammonium acetate (10 mM), apparent pH 7.7

Benzodiazepines (nitrazepam, Cholesteryl bonded silica, nimetazepam, estazolam, 6.5 gm, 300 A brotizolam, clotiazepam, oxazolam, haloxazolam, cloxazolam, medazepam) Benzodiazepines (nitrazepam, Cholesteryl modified etched clotiazepam, cloxazolam, capillary medazepam)

305 mm x 100 gm i.d.

220 mm active length, chiral separation

175

Acetonitrile-5 mM Tris.HC1 200 mm (effective buffer, pH 7.3 (35:65 or 25:75) length) x 100 gm i.d.

Comparison with ODS packing

176

Acetonitrile-10 mMTris.HC1 buffer, pH 7.3 (10-20:90-80)

75 gm i.d.

Open tubular CEC, comparison with packed capillary CEC

176

250 or 400 mm packed length

177

t~

Plant constituents

Cannabinoids (cannabigerol, cannabidiol, cannabinol, A-9-tetrahydrocannabinol, A-8-tetrahydrocannabinol, cannabichromene, A-9-tetrahydrocannabinolic acid)

Hypersil C 18 or Hypersil C8, 3 gm

6.25-25 mM phosphate, pH 2.57 (67-75:35-25)

340 or 490 mm x 100 gm i.d.

Morphine alkaloids

Nucleosil 100-C 18, 5 gm

Acetonitrile-2 mM ammonium acetate (40:60)

150 mm x 220 ~tm i.d.

118

TABLE 10.1 (continued) bO

Stationary phase

Mobile phase


Note

Ref.

Iodide, iodate and perrhenate

Nucleosil SB, 5 gm

5 mMPhosphate buffer, pH2.6

600 mm x 75 ~tm i.d.


178

Alkali metals and ammonium

Fused silica capillary 5 mMimidazole-10 mMMES 370 mm x 75 ~tm i.d. suspension of cation exchange [2-(N-morpholino)ethaneparticles (slid-phase reagent, sulfonic acid], pH 6.15 SPR)

305 mm effective length, ion-exchange electrochromatography

179

Compound

Inorganic compounds

Applications

413

10.16 REFERENCES

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Index of Compounds Separated A Acebutolol 407 Acenaphthene 344, 373,375,377 Acenaphthylene 344, 372, 373,375 Acetaldehyde 379 Acetaminophen 80, 403,409 Acetone 347, 372, 373,375, 379 Acetophenone 187, 347, 378, 379 Acetylaminofluorene deoxyguanosine adduct 311, 312 Acetylsalicylic acid 80, 320, 381,403 Acidic analytes 96, 97, 100, 105, 131 drugs 364, 401 Acids 98, 100, 104 see also individual acids Adenine 259, 363,396 Adenosine 259, 362, 363,395,396 Adenosine-styrene oxide adducts 395 ADP 126, 396 [3-Adrenergetic blocking agents 331, 410 Adrenosterone 385,386 Alanine and derivatives 264, 276, 299-301, 303,305,359 Albumin, tryptic digest 392 Aldehydes 60 Aldopentoses 380 Aldosterone 124, 306, 385, 386 Aliphatic hydrocarbons 343 Alkali metals 412 Alkaloids 397 see also individual alkaloids Alkylbenzenes 173, 174, 177, 212, 372, 374-376 see also individual alkylbenzenes Alkyl benzoates 382 Alkylphenones 187, 379 see also individual alkylphenones Alkyl phenoxypropanoates, chlorinated 383,408

Alprenolol 407 Ambroxol 75, 410 Ambroxolol 367 Amines 104, 388,403 Aminoglutethimide 330 Amino acids 100, 283,333,358-360, 389-391 acetyl 391 benzoyl 391 benzyloxycarbonyl 391 dansyl derivatives 389, 391 dinitrobenzoyl methyl esters 264 N-(3,5-dinitrobenzyloxycarbonyl) derivatives 391 2,4-dinitrophenyl derivatives 389, 391 enantiomers 389-391 9-fluorenylmethoxycarbonyl derivatives 391 phenylthiohydantoin (PTH) derivatives 80, 276, 299-301,303-305, 358, 359, 389, 391 see also individual amino acid derivatives Aminoglutethimide 407, 411 2-Amino-3-hydroxy-3-phenylpropanol 404 2-Aminopyridine 219 Amitriptyline 128, 199, 410 Ammonium 412 AMP 126, 396 Amphetamine 364, 365,400, 401 Amylase 394 Amylbenzene 157, 215,220, 376 Amylobarbital 123, 401 Amylparaben 348 Androstan-3,17-dione 356, 357, 386 5-c~-Androstan-17-one 204 Androsten-3,17-dione 356, 357, 386 Androstenedione 123,385 Androsterone 124, 204, 205 Angiotensins 209, 260, 392, 393 Anhydrotetracycline 370

422 Anilines 212, 219 Anions, see individual anions; Acidic analytes Anthracene 174, 192, 196, 344, 372, 373, 375-377 Anthraquinone dyes 124 1-(9-Anthryl)-2,2,2-trifluoro-ethanol 407 Antibiotics 258, 368-370, 397 see also individual antibiotics Anti-depressants 121, 122, 126, 128, 199 see also individual anti-depressants Anti-epileptics 126, 404 see also individual anti-epileptics Anti-inflammatory drugs, non-steroidal 73, 403,410 Antipyrine 75,367, 410 Antiviral drugs 126, 403 see also individual antiviral drugs Aromatic acids 219 Aromatic compounds 97, 196 Aromatic hydrocarbons 60 Asparagine and derivatives 276, 299-301, 303-305,359 Aspartame 258, 395 Atenolol 407 ATP 126, 396 Atrazine 399 Azo dyes 124 B

Barbital 123,401 Barbiturates 96, 97, 123,402, 405,408 see also individual barbiturates Barbituric acids 327 alkylated 406 Bases 98, 104 see also individual basic compounds Basic analytes 96, 97, 105,267, 331 drugs 364, 401 peptides 394 proteins 394 Bendrofluazide 123,402 Bendroflumethiazide 125,402 Benz[a]anthracene 349 Benzaldehyde 215,220, 236, 237, 376, 377,379 Benzamide 91 Benz[a]anthracene 373,374

Index

1,2-Benzanthracene 372 Benzene 115,215,220, 344, 346, 373 derivatives 215,374-377, 388 Benzoate derivatives 76, 79 Benzo[g]chrysene adduct with deoxyguanosine 311, 312 Benzodiazepines 124, 263,264, 307, 368, 408,409, 411 see also individual azepines Benzo[a]pyrene 174, 175,344, 373 Benzo[b]fluoranthene 344, 373 Benzo [g]chrysene- 11,12-dihydrodiol- 13,14epoxide-DNA adducts 59, 60 B e n z o [ g h i ] p e r y l e n e 344, 373 Benzo[k]fluoranthene 344, 373 Benzoic acid 99, 131, 219 derivatives 384, 385 Benzoin 327, 330, 405,407, 408, 410 acetate 333,408 Benzophenone 376, 377, 379 Benzyl alcohol 91,157, 189, 215,220, 236, 237, 376, 377, 379, 400 Benzylamine 90, 92, 93, 99 Berberine 403 Beta-estradiol 124 Betamethasone 124, 386, 387 dipropionate 124, 387 17-valerate 387, 388,400 Bile acids 201,206, 207, 357 1,1 '- Binaphthy1-2,2'-dihydrogenphosphate 406 1,1 '-Binaphthyl-2,2'-diol 408 Biotin 342 Biphenyl 67, 90, 91,157, 321,377, 379, 384, 385 1-(p-Biphenyl)-ethanol 406 Bradykinin 260, 392, 393 Bromides 128 o-Bromobenzoic acid 131 p-Bromobenzoic acid 131 4-Bromobenzoic acid 219 Brotizolam 411 Budesonide 125 Budesonine 387 Bufadienolide steroids 308 Bufalin 125,386 Bumetanide 402, 403 Bupivacaine 333

423

Subject Index

2-Butanone 379 Butethal 123, 401 N-Butylaniline 219 Butylbenzene 115, 215,220, 346, 372, 374, 377 Butylbenzoic acid 324 Butyl-p-hydroxybenzoic acid 384 Butylparaben 348, 372, 376, 377 Butyraldehyde 379 Butyrophenone 187, 347, 378, 379

Caffeine 80, 93, 99, 258,403,409 Cannabichromene 411 Cannabidiol 411 Cannabigerol 411 Cannabinoids 97, 123,364, 411 see also individual cannabinoids Cannabinol 400, 401, 411 Carbamazepine 404 Carbamazepine- 10,11-diol 404 Carbamazepine- 10,11-epoxide 404 Carbohydrates 348-350 see also individual carbohydrates Carbonic anhydrase 394 Carboxylic acids 320, 381-385 see also individual carboxylic acids Carbovir 410 Cardenolide steroids 308 Cardiac glycosides 404 Carprofen 406 Catechol 196, 378 Cefuroxime axetil 125, 126, 307, 397 Cephalosporin 125, 126, 307, 397 Cerbetocin 394 Chlomipramine 402 Chlorides 128 6-Chloro-a-methylcarbazole-2-acetic cid 406 Chlorobenzenes 212 4-Chlorobenzoic acid 219 Chlorocresol 400 Chlorodiazepam 263 O,L-Chlorodiazepoxide 263,405 Chlorotetracycline 370 Chlorthalidone 335,402, 405 Cholic acid 204 Chrysene 373

Chymotrypsinogen 194, 211,394 Cicloprofen 406 Cinobufagin 125,386 Cinobufatalin 125, 386 Cinosulfuron 369, 371,398 by-products 369, 371,398 Clenbuterol 405,409 Clobetasol butyrate 124, 387 Clobetasol- 17-propionate 400 Clobetasone butyrate 387 Clomipramine 405, 410 Clonazepam 409 Clotiazepam 403, 411 Cloxazolam 403, 411 Cocaine 364, 365,400, 401,403 Codeine phosphate 403 1,3,5-Collidine 219 Conalbumin 100 Corticosteroids 386 see also individual corticosteroids Corticosterone 357, 386 Cortisone 386 acetate 400 Coumarin dyes 399 see also individual categories of dyes Cresols 378 Crotonaldehyde 379 Cyanazine 399 Cyclophophamide 407 Cytidine 362, 363,395,396 Cytochrome c 194, 211, 261,262, 361, 393,395 tryptic digest 72, 309, 310, 321,322, 392 Cytosine 259, 363,396

De-esterified steroids 387 Dehydroisoandrosterone 204, 205 Desethylatrazine 399 Deoxyribonucleic acids, see DNA Desipramine 402 Desisopropylatrazine 399 Desmopressin 394 Detamethasone 127 Dexamethasone 124, 127, 385,386, 388, 400

424

2,2'-Diamino-6,6'-dimethylbiphenyl 330, 411 Diazepam 124, 263,365,401,403,405, 409 Dibenz[a,h]anthracene 344, 373 Dibucaine 384 1,4-Dichlorobenzene 376 Diclofop methyl 327, 408 Diethylphthalate 67, 321,384, 385 Digitalis glycosides 357 Digitoxigenin 125,386, 404 Digoxigenin 125,386, 404 Digoxin 404 2',5'-Dihydroxyacetophenone 378 3,5-Dihydroxybenzoic acid 219 3,4-Dihydroxyphenylalanine (DOPA) 404 2',5'-Dihydroxypropiophenone 378 Dimethylnaphthalene 157 Dimethylphthalate 67, 321,384, 385 3,5-Dinitrobenzoic acid 131 N-(3,5-Dinitrobenzoyl)leucine diallylamide enantiomers 227 N-(3,5-Dinitrobenzyloxycarbonyl) amino acids 391 2,4-Dinitrophenyl amino acids derivatives 358 2,4-Dinitrotoluene 388 Dinucleotides 100 see also Nucleotides Diphenhydramine 90, 92 Disopyramide 407 Diuretics 60, 123,125,307 see also individual diuretics DNA adducts 59, 60, 311 DNB-alanine 405 DNB-leucine 228,229, 331,391 DNZ-amino acids 390 DNZ-leucine 230, 391 DOPA, s e e 3,4-Dihydroxyphenylalanine Doxepine 405 Drugs 235,258-260, 266, 363-368, 400chiral 263,264 metabolites 258 of abuse 364, 400 Dyes 399

Index

E

Econazole 330, 411 Eicosenoic acid 384 Enkephalines 216, 392, 393 Ephedrine 403,404 4-Epianhydrotetracycline 370 Epinephrine 404 4-Epitetracycline 370 Epitizide 123,402 Equiline 204 Erucic acid 384 Estazolam 411 Estradiol 302, 304, 385,387 Estriol 387 Estrone 204, 205,387 Ethosuccinimide 126, 404 N-Ethylaniline 219 Ethylbenzene 115,215,220, 346, 372, 374,376,377 5-Ethyl- 1-methyl-5-(n-propyl) barbituric acid 405 Ethylnaphthalene 157 Ethylparaben 348, 372, 376, 377 Etodolac 406, 410 Etozolin 330, 411 Explosives 119 F

FAPEs, s e e Fatty acids, phenacyl esters Fatty acids 125,350-354, 383,384 methyl esters 302, 350, 384 phenacyl esters 350, 351, 353, 354, 383 Fenoxaprop methyl 327, 408 Fluocortolone 124, 385,386 Fluoranthene 119, 344, 372, 373,375-377 Fluorene 174, 192, 344, 349, 372-377 Fluorides 128 p-Fluorophenylalanine 390 Flurbiprofen 403,406 Fluticasone 385 propionate 127, 387, 388 Fluvoxamine 388 Fmoc-amino acids 390 Fmoc-leucine 331,332, 391 Folic acid 320, 381 Food dyes 127, 399

425

Subject I n d e x

Formaldehyde 379 Formamide 356, 357, 359 Formates 128 Fullerenes 374 G Gitoxigen 386 Glucose 348, 350 Glucuronides, aromatic 380 Glutamine and derivatives 276, 299-301, 303,305,359 Gluthethimide 327, 330, 405, 411 Glycine and derivatives 276, 299-301,303, 305, 359 conjugates 204, 207 Glycosphingolipids 381 Growth hormone, tryptic digest 395 Guanine 259, 363,396 Guanosine 362, 363,395, 396 H

Halogenated hydrocarbons 345 Haloxazolam 411 Herbicides 399 Heroin 364, 365,400, 401 Hexaldehyde 379 Hexanophenone 187 Hexobarbital 121,123,331,401,405-408 Hexylbenzene 175, 176 Hexylparaben 372, 376, 377 Homoatropine 405 Hydrocarbons 343-348, 372-377 Hydrochlorothiazide 409 Hydrocortisone 124, 306, 385-387, 400 derivatives 387 Hydrocortisone- 17-butyrate 400 Hydrocortisone- 17-valerate 400 Hydroflumethiazide 402 Hydrophobic compounds 320 Hydroquinone 196, 379 ethers 196, 379 1113-Hydroxyandrosterone 204, 205 19-Hydroxy-4-androsterone-3,17-dione 204 2-Hydroxyatrazine 399 p-Hydroxybenzoic acid 93,320, 381,384 4-Hydroxybenzoic acid 219 p-Hydroxybenzoic acid 403

c~-Hydroxyethyl-naphthalene 330 13-Hydroxyethyltheophylline 409 17-ct-Hydroxyprogesterone 385 20-c~-Hydroxyprogesterone 385 Hydroxyquinone 378 2-Hydroxyterbutylazine 399 I

Ibuprofen 403,406 IgG, see Immunoglobulin G Imipramine 402, 410 Immunoglobulin G 342 Indapamide 330, 410, 411 Indeno[1,2,3-c,d]pyrene 344, 373 Indomethacin 403 Inorganic compounds 100, 128, 412 see also individual inorganic compounds Inosine 362, 363,396 Inosine-styrene oxide adducts 395 Insecticides 346 Insulin 209, 395 Iodate 99, 412 Iodide 99, 412 4-Iodobenzoic acid 219 Isofamide 407 Isoleucine and derivatives 299-301,358, 359 Isradepin 124, 404 by-products 124

Jatrorrizine 403 K

Ketones 60, 347, 348 Ketoprofen 403

a-Lactalbumin 395 [3-Lactoglobulin 392 Lanthanides 100 Leucine and derivatives 228-230, 299-301, 331,358, 359, 390, 391 see also Amino acids Leucine enkephaline 216

426 Lipids 353 Lorazepam 330, 410 Lysozyme 194, 211, 261,264, 265, 361, 393,395 M

Macrocyclic antibiotics 397 Macrocyclic lactones 125 Malonic acid, sulfate 382 Maltohexaose 348, 350 Malto-oligosaccharides 348-350, 352, 380 Mecoprop methyl 327, 408 Medazepam 411 Mephobarbital 120, 327-329, 405,407, 408 Mepivacaine 333 Methacrolein 379 Methadone 409 Methamphetamine 364, 365,400, 401 Methaqualone 365,366, 401 Methdilazine 402 Methionine enkephaline 216 Methyl amitriptyline 199 Methyl benzoate 376 Methylbenzoin 333,408 Methylchlothiazide 402 Methyl-p-hydroxybenzoic acid 384 Methyl-mandelate 327, 405 Methylnaphthalene 192, 236, 237, 375 Methylparaben 189, 348 c~-Methyl-2,3,4,5,6-pentafluorobenzyl alcohol 406 a-Methyl-a-phenyl-succinimide 327, 405 Methylprednisolone 386 1-Methyl-5-(2-propyl)-5-(n-propyl)barbituric acid 405 17-a-Methyltestosterone 387 Methylthiohydantoin-proline 405 Metoclopramide 75,367, 410 Metolazone 123,402 Metomidate 330, 411 Metoprolol 333,407 Mianserin 406 Monosaccharides 381 see also individual carbohydrates Morphine alkaloids 126, 397, 411 MTH-proline 327, 408 Myoglobin 261,361,393-395

Index

N

Naphthalene 119, 174, 192, 344, 349, 372377 [3-Naphthol 189, 196 1-(2-Naphthyl)ethanol 327, 406 Naproxene 75,367, 403, 410 Neurotensin 392 Neutral analytes 126 compounds 104, 128 drugs 364 Nicotinamide 320, 381 Nicotinic acid 320, 381 Nimetazepam 411 Nitrated hydrocarbons 344, 345 Nitrazepam 124, 403,409, 411 Nitro compounds 388 Nitrobenzene 375,388 p-Nitrobenzoic acid 131 Nitrogen-containing heterocycles 345 p-Nitrophenyl-~-D-glucopyranosides 349, 352, 380 p-Nitrophenyl-ot-o-maltoside 352, 380 p-Nitrophenyl-a-D-maltopentaoside 352, 380 p-Nitrophenyl-ot-D-maltotetraoside 352, 380 p-Nitrophenyl-a-D-maltotrioside 352, 380 p-Nitrophenylglycosides 349 Nitroso compounds 388 N-Nitrosodiethanolamine 388 Nonapeptides 392 see also Peptides Non-ionised (non-polar) analytes 104, 302 Norethindrone 123,385 Nortriptyline 90, 92, 128, 199, 402 Noscapine 364, 365,400, 401 Nucleic acids 100, 311, 312 see also DNA; RNA Nucleobases 100, 259, 363,396 Nucleosides 97, 125,362, 363,395,396 Nucleotides 126, 128, 396 O Octanophenone 187 Oils, vegetable 125 Oleic acid 384

Subject Index

Oligonucleotides 277 see also Nucleotides Oligopeptides 201, 210 see also Peptides Oligosaccharides 201,202, 348, 351,380 see also individual carbohydrates Ondansetron 410 Oxazepam 263,409 Oxazolam 411 Oxygen-containing heterocycles 345 Oxytocin 394 Ovalbumin, tryptic digest 394 P

PAHs, see Polyaromatic hydrocarbons Palmitoleic acid 384 Parabens 127, 348,372, 376, 400 Paracetamol, metabolites 80 Pentapeptides 392 Pentobarbital 405,407 Pentylbenzene 115,346, 372, 374 Pentylparaben 372, 376, 377 Peptide drugs, metabolites 68 Peptides 72, 73, 125, 131,132, 201,209, 210, 216, 260-262, 277, 309, 310, 321, 322, 359-362, 391-395 C-protected 393 N-protected 393 Perchlorates 128 Perrhenate 412 Pesticides 346, 369, 398, 399 Pharmaceuticals 97, 104, 258-260 diverse 65 Phenanthrene 174, 192, 196, 344, 349, 372377 Phenobarbital 123,365,366, 400, 401 Phenols 97, 196, 212, 372, 378, 388 Phenothiazine 376 Phenylalanine and derivatives 299, 300, 334, 358, 359, 389, 390 amide 390 Phenylbutazone 409 T-Phenyl-T-butyrolactone 327 Phenylenediamines 212 1-Phenylethanol 406 2-Phenylethylamine derivatives 404 Phenylglycine 390 2-Phenylmethyl-1-naphthol 378

427 Phenyl propanol 388,406 2-Phenylpropionic acid 382 Phenylthiohydantoin-amino acids 80, 276, 299-301,303-305,358, 359, 389, 391 Phenytoin 126, 404 Phthalate esters 382 Pindolol 407 Piprozolin 330, 411 Polyaromatic hydrocarbons (PAHs) 59, 78, 116, 127, 173, 174, 177, 343-345,372377, 379, 409 see also individual polyaromatic hydrocarbons Polychlorinated dibenzo-p-dioxins 399 Polystyrene standards 224, 376 Prednisolone 124, 386, 387 acetate 400 Pregnan-3,20-dione 356, 357, 386 4-Pregnen-20~-ol-3-one 387 Prenalterol 407 Preservatives 97 Primicarb 398 Primidon 126, 404 Procainamide 90-92 Procaine 75,364, 365,367, 400, 401, 410 Profens 391 Progesterone 123,204, 356, 385,387 metabolites 356 Proline and derivatives 299-301,327, 359 Propazine 399 Propionaldehyde 379 Propiophenone 378 Propranolol 333,384, 407, 408 Propylbenzene 115, 215,220, 346, 372, 374,376 Propyl-p-hydroxybenzoic acid 384 Propylparaben 348, 372, 376, 377 Prostaglandins 126, 381 Protein(s) 100, 194, 201, 211,225,260262, 264, 309, 361,392-395 digests 309, 310, 361,392 sequencing 283 see also individual proteins PTH-amino acids, see Phenylthiohydantoinamino acids Purine bases 128, 363,396 see also Nucleobases Pyrene 174, 196, 344, 349, 373-375,377

428 9-(1-Pyrene)nonanol 137 Pyridine 378 Pyrimidine bases 128, 363,396 see also Nucleobases

Q Quinine 364, 365,400, 401 R

Ranitidine 410 Resorcinol 189, 196, 378 Retinyl esters 123,383,397 Ribonucleases 194, 211, 261,264, 361, 393-395 t-RNA 100, 128, 312 Ropivacaine 333 S

S-oxidation compounds 124 Salbutamol 409 Salsolinol 335,406 Sebutylazine 399 Secobarbital 123, 401 Serin 390 Serotonin 258,265 Simazine 399 Softeners 346 Sorbic acid 400 Steroid A 125 Steroid drugs 355 glycosides 357 hormones 357 Steroids 124, 125, 188, 201,203-205,307, 308,355-358,385-388 see also individual steroids trans-Stilbene oxide 330, 333,408, 410, 411 Sugars 201 Sulfadicramide 404 Sulfaflurazol 404 Sulfanilamide 404 Sulfides 128 Sulfonates, aliphatic 128 Sulphur-containing heterocycles 345 Suramin 126 Sweeteners, artificial, see Aspartame

lndex

T Taurine conjugates 204, 207 Temazepam 263,405,407, 409 Terbutaline 408 Terbutylazine 399 o-Terphenyl 67, 321,384, 385 Testosterone 123, 124, 306, 357, 365, 385387, 400, 401 decanoate 383 esters 353 isocaproate 383 phenyl propionate 383 propionate 365,383,400, 401 1,2,4,5-Tetrachlorobenzene 376 Tetracyclines 258, 370, 397 A-8-Tetrahydrocannabinol 411 see also Cannabinoids A-9-Tetrahydrocannabinol 365,400, 401, 411 see also Cannabinoids A-9-Tetrahydrocannabinolic acid 365,400, 401, 411 see also Cannabinoids Tetrapeptides 392, 393 see also Peptides Textile dyes 124, 399 Thalidomide 330-332, 409 metabolites 330, 409 Thebaine 403 Theobromine 409 Theophylline 258, 409 Thiazide diuretics 307 Thiocyanates 128 Thiourea 67, 75, 90, 91, 119, 157, 175, 176, 215,220, 236, 237, 320, 321, 362, 367, 376, 377, 379, 381,384, 385, 388, 400, 410 Thomapyrin 80, 403 Threonine and derivatives 276, 299-301, 303,305,359 Thymidine 362, 395,396 Thymine 259, 363,396 Timolol 75,367, 410 Tipredane 125,369, 385 p-Tolualdehyde 379 o-Toluic acid 131 Toluene 115,215,220, 346, 372, 374, 376

429

Subject Index

2-Toluic acid 219 Triamcinolone 386 acetonide 386 Triazine herbicides 123 see also individual herbicides 1,3,5-Trichlorobenzene 376 Tricyclic anti-depressants 126, 410 see also individual anti-depressants Triglycerides 125,352-355,382, 383 1,2,3-Trimethylbenzene 375 Triphenylene 174 Tripredane 401 Tr6ger's base 330, 333,408,411 Tropicamide 405 Trypsinogen 394 Tryptamine 258,265 Tryptophan and derivatives 299, 300, 331, 359, 390, 391 Tyrosine and derivatives 276, 299-301, 303,305,359, 390

Tyrosine-containing peptides 392 see also Peptides

Uracil 363,396 see also Nucleobases Urea herbicides 398 see also individual herbicides Uridine 362, 363,395,396 see also Nucleosides V Valeraldehyde 379 Valine and derivatives 264, 299-301,358 see also Amino acids Vitamins 302, 397 W Warfarin 330, 406, 407


431

JOURNAL

OF C H R O M A T O G R A P H Y

LIBRARY

A Series of Books Devoted to Chromatographic and Electrophoretic Techniques and their Applications Although complementary to the Journal of Chromatography, each volume in the Library Series is an important and independent contribution in the field of chromatography and electrophoresis. The library contains no material reprinted from the journal itself.

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Chromatography of Antibiotics (see also Volume 26) by G.H. Wagman and M.J. Weinstein

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Extraction Chromatography edited by T. Braun and G. Ghersini

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Liquid Column Chromatography. A Survey of Modern Techniques and Applications edited by Z. Deyl, K. Macek and J. Jan~ik

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Detectors of Gas Chromatography by J. t~ev~ik

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Instrumental Liquid Chromatography. A Practical Manual on HighPerformance Liquid Chromatographic Methods (see also Volume 27) by N.A. Parris

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Isotachophoresis. Theory, Instrumentation and Applications by F.M. Everaerts, J.L. Beckers and Th.P.E.M. Verheggen Chemical Derivatization in Liquid Chromatography by J.F. Lawrence and R.W. Frei

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Chromatography of Steroids by E. Heftmann

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HPTLC - High Performance Thin-Layer Chromatography edited by A. Zlatkis and R.E. Kaiser

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Gas Chromatography of Polymers by V.G. Berezkin, V.R. Alishoyev and I.B. Nemirovskaya

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Liquid Chromatography Detectors

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Affinity Chromatography (see also Volume 55)

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Radiochromatography. The Chromatography and Electrophoresis of Radiolabelled Compounds by T.R. Roberts

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432

Volume 16

P o r o u s Silica. Its Properties and Use as Support in Column Liquid Chromatography by K.K. Unger

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75 Years of C h r o m a t o g r a p h y - A Historical Dialogue edited by L.S. Ettre and A. Zlatkis

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E l e c t r o p h o r e s i s . A Survey of Techniques and Applications P a r t A: T e c h n i q u e s P a r t B: Applications edited by Z. Deyl

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C h r o m a t o g r a p h y . Fundamentals and Applications of Chromatographic and Electrophoretic Methods (see also Volume 51) P a r t A: F u n d a m e n t a l s

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C h r o m a t o g r a p h y of Alkaloids P a r t A: T h i n - L a y e r C h r o m a t o g r a p h y by A. Baerheim-Svendsen and R. Verpoorte P a r t B: Gas-Liquid C h r o m a t o g r a p h y a n d H i g h - P e r f o r m a n c e Liquid C h r o m a t o g r a p h y by R. Verpoorte and A. Baerheim-Svendsen

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C h r o m a t o g r a p h y on Antibiotics. Second, Completely R e v i s e d Edition by G.H. Wagman and M.J. Weinstein

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433

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The Science of C h r o m a t o g r a p h y . Lectures Presented at the A.J.P. Martin Honorary Symposium, Urbino, May 27-31, 1985 edited by F. Bruner

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Liquid C h r o m a t o g r a p h y Detectors. Second, C o m p l e t e l y R e v i s e d Edition by R.P.W. Scott

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434

Volume 45A

C h r o m a t o g r a p h y and Modification of N u c l e o s i d e s Analytical Methods for Major and Modified N u c l e o s i d e s HPLC, GC, MS, NMR, UV and FT-IR edited by C.W. Gehrke and K.C.T. Kuo

Volume 45B

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s Biological Roles a n d F u n c t i o n of Modification edited by C.W. Gehrke and K.C.T. Kuo

Volume 45C

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s Modified N u c l e o s i d e s in C a n c e r a n d Normal Metabolism Methods and Applications edited by C.W. Gehrke and K.C.T. Kuo

Volume 45D

C h r o m a t o g r a p h y a n d Modification of N u c l e o s i d e s C o m p r e h e n s i v e D a t a b a s e for RNZ and DNA N u c l e o s i d e s Chemical, Biochemical, Physical, Spectral and S e q u e n c e edited by C.W. Gehrke and K.C.T. Kuo

Volume 46

Ion C h r o m a t o g r a p h y : Principles and Applications by P.R. Haddad and P.E. Jackson

Volume 47

Trace Metal Analysis and Speciation edited by I.S. Krull

Volume 48

S t a t i o n a r y P h a s e s in Gas C h r o m a t o g r a p h y by H. Rotzsche

Volume 49

Gas C h r o m a t o g r a p h y in Air Pollution Analysis by V.G. Berezkin and Yu.S. Drugov

Volume 50

Liquid C h r o m a t o g r a p h y in B i o m e d i c a l Analysis edited by T. Hanai

Volume 51

C h r o m a t o g r a p h y , 5th edition. Fundamentals and Applications of Chromatographic and Related Differential Migration Methods Part A: F u n d a m e n t a l s and Techniques P a r t B: Applications edited by E. Heftmann

Volume 52

Capillary E l e c t r o p h o r e s i s . Principles, Practice and Applications by S.F.Y. Li

Volume 53

H y p h e n a t e d Techniques in Supercritical Fluid C h r o m a t o g r a p h ~ and Extraction edited by K. Jinno

Volume 54

C h r o m a t o g r a p h y of Mycotoxins. Techniques and Applications edited by V. Betina

Volume 55

Bioaffinity C h r o m a t o g r a p h y . Second, completely revised edition by J. Turkov~

Volume 56

C h r o m a t o g r a p h y in the P e t r o l e u m I n d u s t r y edited by E.R. Adlard

Volume 57

Retention and Selectivity in Liquid C h r o m a t o g r a p h y . Prediction, Standardisation and Phase Comparisons edited by R.M. Smith

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Volume 58

C a r b o h y d r a t e Analysis edited by Z. E1 Rassi

Volume 59

Applications of Liquid C h r o m a t o g r a p h y / Mass S p e c t r o m e t r y in Environmental Chemistry edited by D. Barcel5

Volume 60

A d v a n c e d C h r o m a t o g r a p h i c a n d E l e c t r o m i g r a t i o n M e t h o d s in BioSciences edited by Z. Deyl, I. Mikgik, F. Tagliaro and E. Tesa~ov~

Volume 61

P r o t e i n Liquid C h r o m a t o g r a p h y edited by M. Kastner

Volume 62

Capillary E l e c t r o c h r o m a t o g r a p h y edited by Z. Deyl and F. Svec


Capillary Electrochromatography

Capillary Electrochromatography

Capillary Electrochromatography (RSC Chromatography Monographs)

Capillary Electrophoresis

Capillary Electrophoresis

Capillary electrophoresis of carbohydrates

Capillary Electrophoresis of Nucleic Acids. Practical Applications of Capillary Electrophoresis

Capillary Electrophoresis Technology

Capillary Electrophoresis Technology

Practical Capillary Electrophoresis

capillary gas adsorption chromatography

Capillary Electrophoresis of Carbohydrates

Capillary Gas Adsorption Chromatography

Capillary Condensation and Adsorption

Capillary Electrophoresis of Nucleic Acids. Introduction to the Capillary Electrophoresis

Capillary Electrophoresis of Nucleic Acids. Practical Applications of Capillary Electrophoresis

Capillary Flows with Forming Interfaces

Capillary Electrophoresis in Chiral Analysis

Capillary Flows with Forming Interfaces

Capillary Electrophoresis. Methods and Protocols

Capillary Electrophoresis: Methods and Protocols

Capillary Electrophoresis in Polymer Analysis

Clinical Applications of Capillary Electrophoresis

Practical Capillary Electrophoresis, Second Edition

Gravity-Capillary Free-Surface Flows

Chemometric Methods in Capillary Electrophoresis

Capillary Electrophoresis Guidebook: Principles, Operation, and Applications

Capillary Electrophoresis of Proteins and Peptides

Chiral Separations by Capillary Electrophoresis (Chromatographic Science)

Capillary Fluid Exchange: Regulation, Functions, and Pathology

Analysis and Detection by Capillary Electrophoresis

Capillary Electrochromatography