analytical separations using packed and open-tubular
TRANSCRIPT
Louisiana State UniversityLSU Digital Commons
LSU Doctoral Dissertations Graduate School
2004
Analytical separations using packed and open-tubular capillary electrochromatographyConstantina P. KapnissiLouisiana State University and Agricultural and Mechanical College, [email protected]
Follow this and additional works at: https://digitalcommons.lsu.edu/gradschool_dissertations
Part of the Chemistry Commons
This Dissertation is brought to you for free and open access by the Graduate School at LSU Digital Commons. It has been accepted for inclusion inLSU Doctoral Dissertations by an authorized graduate school editor of LSU Digital Commons. For more information, please [email protected].
Recommended CitationKapnissi, Constantina P., "Analytical separations using packed and open-tubular capillary electrochromatography" (2004). LSUDoctoral Dissertations. 595.https://digitalcommons.lsu.edu/gradschool_dissertations/595
ANALYTICAL SEPARATIONS USING PACKED AND OPEN-TUBULAR CAPILLARY ELECTROCHROMATOGRAPHY
A Dissertation
Submitted to the Graduate Faculty of the
Louisiana State University and Agricultural and Mechanical College
in partial fulfillment of the requirements for the degree of
Doctor of Philosophy
in
The Department of Chemistry
by Constantina P. Kapnissi
B.S., University of Cyprus, 1999 August 2004
ii
Copyright 2004 Constantina Panayioti Kapnissi All rights reserved
iii
DEDICATION
I would like to dedicate this work to my husband Andreas Christodoulou, my parents
Panayiotis and Eleni Kapnissi, and my sisters Erasmia, Panayiota and Stella Kapnissi. I
want to thank all of you for helping me, in your own way, to finally make one of my
dreams come true. Thank you for encouraging me to continue and achieve my goals.
Thank you for your endless love, support, and motivation. Andreas, thank you for your
continuous patience and for being there for me whenever I needed you during this difficult
time. I will never forget what you did for me.
iv
ACKNOWLEDGMENTS
I would like to acknowledge the following people for their contributions in this work:
Dr. Isiah M. Warner for his guidance and excellent advice. Thank you for encouraging
me to finish what I started. I want to also thank you for the financial and moral support.
You are an excellent mentor, and you prove that every day with your constant
encouragement and support. Thank you again for everything.
Dr. Rezik A. Agbaria for his helpful scientific discussions during my general exam and
my project with laser scanning confocal microscopy.
Dr. Charles William Henry III for training me how to use the capillary electrophoresis
instrument and the packing apparatus. Dr. Henry and Dr. Shahab Shamsi helped me
understand the most important concepts of my research.
Dr. Joseph B. Schlenoff and his group at Florida State University in Tallahassee,
Florida for training me on the polyelectrolyte multilayer coating procedure.
Dr. Lei Geng and Mark Lowry at University of Iowa in Iowa City, Iowa for helping
me conduct some experiments on the laser scanning confocal microscope, and for their
helpful discussions about this part of my research.
Dr. Akbay Cevdet and Bertha C. Valle for providing the polymeric surfactants I used in
my research work.
Warner research group and Warner family for their encouragement and their total
support.
Bertha C. Valle for her friendship, her advice, and the long useful discussions we had
about research.
v
Dr. Nikki Gill, Dr. Xiaofeng Zhu, and Dr. Abdul Numan for editing a few of my
reports.
My family and friends in Cyprus for believing in me, and supporting all my choices.
vi
TABLE OF CONTENTS
DEDICATION .....................................................................................................................iii
ACKNOWLEDGMENTS ...................................................................................................iv
LIST OF TABLES ...............................................................................................................ix
LIST OF FIGURES ..............................................................................................................x
LIST OF ABBREVIATIONS ............................................................................................xiv
ABSTRACT .................................................................................................................... xviii
CHAPTER 1. INTRODUCTION .........................................................................................1 1.1 Capillary Electrophoresis ....................................................................................1 1.2 Micellar Electrokinetic Chromatography ...........................................................9 1.3 Capillary Electrochromatography .....................................................................15
1.3.1 Packed-Capillary Electrochromatography ...............................................17 1.3.2 Open-Tubular Capillary Electrochromatography ....................................18
1.3.2.1 Adsorption ...................................................................................20 1.3.2.2 Covalent Bonding and/or Crosslinking .......................................29 1.3.2.3 Porous Layers ..............................................................................31 1.3.2.4 Chemical Bonding After Etching ................................................32 1.3.2.5 Sol-Gel Technique .......................................................................35
1.4 Chirality ............................................................................................................38 1.5 Chiral Selectors in Open-Tubular Capillary Electrochromatography ..............40
1.5.1 Derivatized Cyclodextrins .......................................................................40 1.5.2 Cellulose ..................................................................................................42 1.5.3 Proteins ....................................................................................................43 1.5.4 Molecular Imprinted Polymers ................................................................44 1.5.5 Polymeric Surfactants ..............................................................................46
1.6 Laser Scanning Confocal Microscopy ..............................................................47 1.7 Scope of Dissertation ........................................................................................51 1.8 References .........................................................................................................54
CHAPTER 2. SEPARATION OF BENZODIAZEPINES BY USE OF CAPILLARY ELECTROCHROMATOGRAPHY .....................................65 2.1 Introduction .......................................................................................................65 2.2 Experimental .....................................................................................................68
2.2.1 Reagents and Chemicals ..........................................................................68 2.2.2 Instrumentation and Conditions ...............................................................68 2.2.3 Sample and Buffer Preparation ................................................................69 2.2.4 Preparation and Conditioning of Packed Capillary Columns ..................69
2.3 Results and Discussion .....................................................................................70
vii
2.3.1 Effect of the Nature of Organic Modifier ................................................70 2.3.2 Effect of Mobile-Phase Composition ......................................................73 2.3.3 Effect of Applied Voltage ........................................................................73 2.3.4 Effect of Tris Concentration ....................................................................75 2.3.5 Effect of Column Temperature ................................................................76 2.3.6 CEC Separation of Drugs from a Urine Sample ......................................78
2.4 Conclusion ........................................................................................................81 2.5 References .........................................................................................................81
CHAPTER 3. ANALYTICAL SEPARATIONS USING MOLECULAR MICELLES IN OPEN-TUBULAR CAPILLARY ELECTROCHROMATOGRAPHY .....................................85 3.1 Introduction .......................................................................................................85 3.2 Experimental .....................................................................................................87
3.2.1 Apparatus and Conditions ........................................................................87 3.2.2 Reagents and Chemicals ..........................................................................87 3.2.3 Sample and Buffer Preparation ................................................................88 3.2.4 Synthesis of Monomeric and Polymeric Surfactant ................................88 3.2.5 Procedure for Polyelectrolyte Multilayer (PEM) Coating .......................88
3.3 Results and Discussion .....................................................................................89 3.3.1 Endurance of PEM Coating .....................................................................89 3.3.2 Stability of PEM Coating .........................................................................91 3.3.3 Reproducibilities ......................................................................................92 3.3.4 Voltage Study ..........................................................................................92 3.3.5 Temperature Study ...................................................................................92 3.3.6 Comparison Between Monomeric and Polymeric Surfactants ................94
3.4 Conclusion ........................................................................................................95 3.5 References .........................................................................................................98
CHAPTER 4. CHIRAL SEPARATIONS USING POLYMERIC SURFACTANTS AND POLYELECTROLYTE MULTILAYERS IN OPEN-TUBULAR CAPILLARY ELECTROCHROMATOGRAPHY .................................. 101 4.1 Introduction .................................................................................................... 101 4.2 Experimental .................................................................................................. 103
4.2.1 Apparatus and Conditions ..................................................................... 103 4.2.2 Reagents and Chemicals ....................................................................... 104 4.2.3 Sample and Background Electrolyte Preparation ................................. 104 4.2.4 Synthesis of Polymeric Surfactant ........................................................ 105 4.2.5 Procedure for Polyelectrolyte Multilayer Coating ................................ 106
4.3 Results and Discussion .................................................................................. 106 4.3.1 Effect of Additives in Polymer Deposition Solutions .......................... 107 4.3.2 Effect of NaCl Concentration ............................................................... 107 4.3.3 Effect of Column Temperature ............................................................. 108 4.3.4 Effect of Bilayer Number ..................................................................... 108
viii
4.3.5 Reproducibilities ................................................................................... 113 4.3.6 Chiral Separation of Analytes ............................................................... 113
4.4 Conclusion ..................................................................................................... 118 4.5 References ...................................................................................................... 118
CHAPTER 5. INVESTIGATION OF THE STABILITY OF POLYELECTROLYTE MULTILAYER COATINGS IN OPEN-TUBULAR CAPILLARY ELECTROCHROMATOGRAPHY USING LASER SCANNING CONFOCAL MICROSCOPY ............................................ 121 5.1 Introduction .................................................................................................... 121 5.2 Experimental .................................................................................................. 123
5.2.1 Apparatus and Conditions ..................................................................... 123 5.2.2 Reagents and Chemicals ....................................................................... 124 5.2.3 Sample and Background Electrolyte Preparation ................................. 124 5.2.4 Synthesis of Polymeric Surfactant ........................................................ 125 5.2.5 Procedure for Polyelectrolyte Multilayer Coating ................................ 125
5.3 Results and Discussion .................................................................................. 126 5.3.1 Open-Tubular Capillary Electrochromatography ................................. 126
5.3.1.1 Coating Stability ....................................................................... 126 5.3.1.2 Capillary Recovery and Coating Regeneration ........................ 130
5.3.2 Laser Scanning Confocal Microscopy .................................................. 133 5.4 Conclusion ..................................................................................................... 137 5.5 References ...................................................................................................... 138
CHAPTER 6. CONCLUSIONS AND FUTURE STUDIES .......................................... 141 6.1 References ...................................................................................................... 147
VITA ................................................................................................................................ 148
ix
LIST OF TABLES
Table Page
1.1 Classification of surfactant molecules used in MEKC ...........................................11
3.1 Reproducibilities of PEM capillary coating. Conditions: same as Figure 3.3 ........94
4.1 Run-to-run reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration was varied ...........................................114 4.2 Capillary-to-capillary reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration was varied...............116 4.3 Reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration, 0.1 M.......................................................116 5.1 Stability of PEM capillary coating. Conditions: same as Figure 5.1 ................... 130
x
LIST OF FIGURES
Figure Page
1.1 Schematic diagram of a CE system ........................................................................ 2
1.2 Origin of EOF – Electrical Double Layer .................................................................5
1.3 (a) EOF flow profile, (b) Pressure driven flow profile and their corresponding solute zone profiles ...........................................................................6
1.4 Solute migration ........................................................................................................7
1.5 Structure of a surfactant molecule ..........................................................................10
1.6 Equilibria between a surfactant molecule, a surface monolayer, and a micelle .....12
1.7 Elution time window for neutral analytes in MEKC ..............................................15
1.8 Origin of EOF in a packed capillary .......................................................................19
1.9 (a) Intrinsic, (b) Extrinsic charge compensation .....................................................26
1.10 Scheme of the PEM-coated capillary .....................................................................27
1.11 Model for the aperture-based near-field optical microscope ..................................49
1.12 Schematic diagram of a confocal microscope ........................................................52
2.1 Structures of the seven benzodiazepine analytes ....................................................71
2.2 Effect of the nature of organic modifier on the CEC separation of benzodiazepines. Conditions: C18 stationary phase; 40 cm packed x 100 µm i.d.; electrolyte, 10 mM Tris (pH 8)-ACN (40:60), 10 mM Tris (pH 8)-MeOH (30:70), 10 mM Tris (pH 8)-THF (55:45); applied voltage, 30 kV; electrokinetic injection, 15 kV for 5 s; temperature, 25 °C; UV detection, 220 nm ...............................................74
2.3 Effect of mobile-phase composition on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the mobile phase composition (ACN-Tris) was varied ...........................................................................................75
xi
2.4 Effect of applied voltage on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the applied voltage was varied; electrolyte, 10 mM Tris (pH 8)-ACN (60:40) ........................................................77
2.5 Effect of Tris concentration on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the Tris buffer concentration was varied; electrolyte, Tris (pH 8)-ACN (60:40); applied voltage, 20 kV ..............................78
2.6 Effect of column temperature on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the temperature was varied; electrolyte, 30 mM Tris (pH 8)-ACN (60:40); applied voltage, 20 kV .....................................79
2.7 CEC separation of drugs from a urine sample. (a) Without spiking. Conditions: C18 stationary phase; 40 cm packed x 100 µm i.d.; electrolyte, 30 mM Tris (pH8)-ACN (60:40); applied voltage, 20 kV; electrokinetic injection, 15 kV for 5 s; temperature, 15 °C; UV detection, 220 nm. (b) With spiking. Separation conditions are the same as above, except electrokinetic injection (urine sample), 20 kV for 10 s; electrokinetic injection (2 mg/ml standard oxazepam), 15 kV for 5 s .........................................................80
3.1 Structures of the (a) monomeric SUG and (b) polymeric SUG ..............................90
3.2 Scheme of the PEM-coated capillary .....................................................................91
3.3 Stability studies of PEM coating. Conditions: 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SUG) with 0.2 M NaCl; pressure injection, 0.1 psi for 1 s; electrolyte, 50 mM Na2HPO4 (pH 9.2); applied voltage, 20 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm ...........................93
3.4 Effect of applied voltage on the OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except applied voltage was varied ......................................................................................96
3.5 Effect of temperature on the OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except the temperature was varied ..........................................................................97
xii
3.6 Comparison between monomeric and polymeric surfactants for OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except applied voltage, 30 kV. (a) 0.5% (w/v) poly (L-SUG) (b) 0.5% (w/v) mono (L-SUG) ...............................98
4.1 Structure of poly (L-SULV) ................................................................................. 105
4.2 Chiral separation of BNP. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV); pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); applied voltage, 30 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm. (a) 0.01 M NaCl; (b) 0.01 M 1E-3MI-HFP; (c) 0.01 M 1B-3MI-TFB ................ 109
4.3 Effect of NaCl concentration on the chiral separation of BNP. Conditions: same as Figure 4.2, except NaCl concentration was varied ............. 110
4.4 Effect of column temperature on the chiral separation of BNP. Conditions: same as Figure 4.2, except temperature was varied ......................... 111
4.5 Effect of bilayer number on the chiral separation of BNP. Conditions: same as Figure 4.2, except bilayer number was varied .................... 112
4.6 Illustration of a run-to-run reproducibility for the chiral separation of BNP. Conditions: same as Figure 4.2, except NaCl concentration, 0.05 M ................................................................................. 115
4.7 (a) Chiral separation of BOH. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); detection, 214 nm. Chiral separation of temazepam. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.9 psi for 7 s; background electrolyte, 25 mM Tris and 25 mM Na2B4O7 (pH 8.5); detection, 220 nm. (c) Chiral separation of secobarbital, and (d) Chiral separation of pentobarbital. Conditions for (c) and (d): 3 bilayers; 0.5% (w/v) PDADMAC and 2.5% (w/v) poly (L-SULV) with 0.2 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 300 mM H3BO3 and 30 mM Na2HPO4 (pH 7.2); detection, 214 nm. Other conditions are the same as Figure 4.2 ......................... 117
xiii
5.1 Dependence of electroosmotic mobility on the NaOH flushing time. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); applied voltage, 30 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm ................................................................................................. 128
5.2 Dependence of selectivity on the NaOH flushing time. Conditions: same as Figure 5.1 ............................................................................ 129
5.3 Coating Regeneration - Dependence of electroosmotic mobility on the NaOH flushing time. Conditions: same as Figure 5.1 .............................. 131
5.4 Dependence of selectivity on the NaOH flushing time. Conditions: same as Figure 5.1 ............................................................................ 132
5.5 The z-Section images of a 20-bilayer capillary (left) and a fused-silica capillary (right). Optical slice thickness: 3.3 µm. z-Step size 1.65 µm. Every fourth image is shown. All images are 204 x 512 pixels (460.6 µm x 115.1 µm) .......................................... 134
5.6 z-Section images of a portion of the same 20-bilayer capillary, as in Figure 5.5. Exposure to 1.0 M NaOH. Optical slice thickness: µm. z-Step size 1.65 µm. Every third image is shown. All images are 512 x 512 pixels (65.8 µm x 65.8 µm) .............................................. 135
5.7 LSCM images of different regions within the same 20-bilayer capillary, as in Figure 5.6. Optical slice thickness: 3.3 µm. Both images are 512 x 820 pixels (115.1 µm x 184.4 µm) ................................. 136
5.8 LSCM image of the same 20-bilayer capillary, as in Figure 5.7. Longer exposure to 1.0 M NaOH. Optical slice thickness: 3.3 µm. The image is 512 x 2048 pixels (115.1 µm x 460.6 µm) ........................................................................................ 137
6.1 Structures of the six β-blocker analytes ............................................................... 142
xiv
LIST OF ABBREVIATIONS
Abbreviation Name
AFM atomic force microscopy
BGE background electrolyte
BNP 1,1’-binaphthyl-2,2’-dihydrogenphosphate
BOH 1,1’-bi-2-naphthol
CDCPC cellulose tris(3,5-dichlorophenylcarbamate)
CE capillary electrophoresis
CEC capillary electrochromatography
CGE capillary gel electrophoresis
CIEF capillary isoelectric focusing
CITP capillary isotachophoresis
CMC critical micellar concentration
CTAB cetyltrimethylammonium bromide
CTAC cetyltrimethylammonium chloride
C18-TEOS n-octadecyltriethoxysilane
CZE capillary zone electrophoresis
DAPS N-dodecyl(N,N-dimethyl-3-ammonio-1-propane sulfonate)
Dioxo[13]aneN4 1,4,7,10-tetraazacyclotridecane-11,13,dione
DMPCC 3,5-dimethylphenylcarbamoyl cellulose
DS dextran sulfate
DTAB dodecyltrimethylammonium bromide
ELP extended light path
xv
EOF electroosmotic flow
FESI field-enhanced sample injection
FRET fluorescence resonance energy transfer
GC gas chromatography
H3BO3 boric acid
HCl hydrochloric acid
HPLC high performance liquid chromatography
H2TPFPP 5,10,15,20-tetrakis(penta fluorophenyl)porphyrin
LC liquid chromatography
LODs limits of detection
LSCM laser scanning confocal microscopy
MEKC micellar electrokinetic chromatography
MIPs molecular imprinted polymers
Mono (L-SUG) mono (sodium N-undecenoyl-L-glycinate)
MS mass spectrometry
NA numerical aperture
Na2B4O7 sodium borate
NaCl sodium chloride
Na2HPO4 sodium phosphate
NaOH sodium hydroxide
NIR near-infrared
NSOM near-field scanning optical microscopy
ODS octadecylsilane
xvi
OT-CEC open-tubular capillary electrochromatography
OT-CLC open-tubular capillary liquid chromatography
Packed-CEC packed capillary electrochromatography
PAHs polycyclic aromatic hydrocarbons
PAPS lithium 3’-phosphoadenosine 5’-phosphosulfate
PB polybrene
PDADMAC poly (diallyldimethylammonium chloride)
PEI polyethyleneimine
PEM polyelectrolyte multilayer
pI isoelectric point
PMBC para-methylbenzoyl cellulose
Poly (L-SUG) poly (sodium N-undecanoyl-L-glycinate)
Poly (L-SULV) poly (sodium N-undecanoyl-L-leucylvalinate)
Poly (SUS) poly (sodium undecylenic sulfate)
PSS poly (styrene sulfonate)
PVA poly (vinylamine)
R6G rhodamine 6G
RSD relative standard deviation
SDS sodium dodecyl sulfate
SFC supercritical fluid chromatography
SMIL successive multiple ionic-polymer layer
TEOS n-octadecyltriethoxysilane
THF tetrahydrofuran
xvii
Tris tris(hydroxymethyl)aminomethane
1B-3MI-TFB 1-butyl-3-methylimidazolium tetrafluoroborate
1E-3MI-HFP 1-ethyl-3-methyl-1H-imidazolium hexafluorophosphate
xviii
ABSTRACT
The goal of the research reported in this dissertation is to develop packed and open-tubular
capillary electrochromatographic methods for improved achiral and chiral separations of
various classes of analytes. The first part of this research involves the separation of seven
benzodiazepines by the use of the packed mode of capillary electrochromatography (CEC)
and a 40 cm packed bed of Reliasil 3 µm C18 stationary phase. Optimal conditions were
established by varying the mobile phase, the amount of organic modifier, the buffer
concentration, the applied voltage, and the column temperature. The second part of this
research focuses on the open-tubular mode of CEC and the polyelectrolyte multilayer
(PEM) coating approach. In the first study of this part, poly (diallyldimethylammonium
chloride), PDADMAC, was used as the cationic polymer and poly (sodium N-undecanoyl-
L-glycinate), poly (L-SUG), was used as the anionic polymer for the construction of the
PEM coating. The performance of the modified capillaries as a separation medium was
evaluated by use of seven benzodiazepines as analytes. In the second study, the anionic
polymeric surfactant poly (sodium N-undecanoyl-L-leucylvalinate), poly (L-SULV), was
used as the chiral discriminator for the separations of several drug analytes.
Reproducibility of the PEM coating was evaluated by computing the relative standard
deviation (RSD) values of the electroosmotic flow (EOF). The PEM-coated capillaries
were remarkably robust with excellent reproducibilities and high stabilities against extreme
pH values. The stability of the capillary surface was further investigated after exposure to
NaOH solutions. The structural changes of these coatings were monitored using laser
scanning confocal microscopy (LSCM). These changes were discussed in terms of
separations using open-tubular CEC (OT-CEC). In addition, the electropherograms
xix
obtained from the chiral separation of 1,1’-binaphthyl-2,2’-dihydrogenphosphate (BNP) in
OT-CEC allowed the measurements of both selectivity and electroosmotic mobility
changes after long exposure to NaOH.
1
CHAPTER 1.
INTRODUCTION
1.1 Capillary Electrophoresis
Capillary electrophoresis (CE) is a family of related techniques that use narrow-bore
fused-silica capillaries to perform high efficiency separations of both small and large
molecules. Although CE is originally considered for the analysis of biological
macromolecules, it has also been utilized for the separation of other compounds such as
chiral drugs, vitamins, pesticides, dyes, inorganic ions, organic acids, and surfactants [1,
2]. CE offers a number of advantages when compared with chromatographic techniques:
(1) extremely small amount of sample; (2) high separation efficiency and resolution; (3)
rapid and quantitative separation; (4) automated instrumentation; (5) various modes to vary
selectivity; and (6) simple separation mechanism [2, 3].
As mentioned above, one of the main advantages of CE is the overall simplicity of the
instrumentation. Figure 1.1 illustrates a schematic diagram of a typical CE system. The
basic components of this system are a fused-silica capillary whose ends are placed in
buffer reservoirs, a high-voltage power supply, a UV lamp, a photodiode-array detector, a
sample reservoir, and two electrodes that are used to make electrical contact between the
high-voltage power supply and the capillary. In CE, the separation of analytes is achieved
by replacing the inlet buffer reservoir with a sample reservoir, and by applying either an
electric field or an external pressure. After the sample is loaded onto the capillary, the
sample is replaced by the inlet buffer reservoir, the electric field is applied, the ions in the
sample move along the capillary, and the separation is performed.
2
Sample ReservoirInlet Buffer Reservoir
(-) Electrode
Anode
Outlet Buffer Reservoir
High-Voltage Power Supply
UV-Lamp
Diode-Array Detector
Capillary
(+) Electrode
Cathode
Figure 1.1 Schematic diagram of a CE system.
The utility of CE is also derived from its six different modes of operation. These modes
of CE include capillary zone electrophoresis (CZE), capillary gel electrophoresis (CGE),
capillary isoelectric focusing (CIEF), capillary isotachophoresis (CITP), micellar
electrokinetic chromatography (MEKC), and capillary electrochromatography (CEC). CZE
is the simplest form and the most widely used technique in CE [1, 2]. The separation
principle of CZE is the difference in charge-to-size ratio and the difference in
electrophoretic mobilities of solutes that result in different velocities. Electrophoretic
3
mobility is the mobility of an ion when an electric field is applied across the capillary. The
velocity of an ion is expressed by:
Eeµν = (1.1)
where ν is the ion velocity, eµ is the electrophoretic mobility, and E is the applied
electric field. The electric field is a function of the applied voltage and the capillary length
with units of V/cm. When a constant electric field is applied, ionic species undergo an
electrostatic force, eF :
qEFe = (1.2)
where q is the charge of a particular ion. This force causes ions to accelerate towards the
oppositely charged electrode. In a viscous medium the electrostatic force is balanced by its
frictional force, fF , that slows the mobility of ions. According to Stokes’ law for spherical
particles, the frictional force can be given by Equation 1.3:
νπηrFf 6= (1.3)
where η is the dynamic viscosity of the solution, and r is the radius of the particle or the
ion. During electrophoresis a steady state is achieved, and at this point the forces are equal.
The combination of the above equations yields an equation that describes the
electrophoretic mobility as follows:
rq
e πηµ
6= (1.4)
The electrophoretic mobility is a property of a given ion and medium, and it is a
physical constant for that ion. Equation 1.4 demonstrates that small species with high
charges have high mobilities, whereas large, minimally charged species have low
4
mobilities. In addition, neutral species do not possess an electrophoretic mobility ( 0=q ).
Therefore, they are not separated by use of CZE. Another important characteristic that is
derived from the above equation is that the electrophoretic mobility of ionic species
decreases as the viscosity of the background electrolyte (BGE) increases [1, 5, 6].
A significant factor that causes the movement of both neutral and charged species in
solution is the electroosmotic flow (EOF). The origin of this flow is the electrical double
layer that is formed at the solid-liquid interface, between the bulk liquid and the inner
capillary surface (Figure 1.2). Under alkaline conditions, the inside wall of a fused-silica
capillary is negatively charged due to ionization of the surface silanol groups to negatively
charged silanoate groups. Electrolyte cations accumulate adjacent to the negative surface
of the capillary, they absorb on it by electrostatic attraction, and they form an immobilized
layer that is called the Stern layer. The remaining ions constitute the diffuse layer, which
extends into the bulk liquid. This arrangement of ions develops the electric double layer.
When an electric field is applied across the column, the cations, which predominate in the
diffuse layer, migrate in the direction of the cathode. Since ions are solvated by water, the
bulk solution in the capillary is dragged along by the migrating charge.
This generates the EOF, which can be described as follows:
EEOF
=ηεζν (1.5)
where EOFν is the electroosmotic velocity, ζ is the zeta potential, and ε is the dielectric
constant of the BGE. The zeta potential is the corresponding potential across the layers,
and it depends on the thickness of the diffuse layer and the surface charge on the capillary
wall. The EOF can be also expressed in terms of mobility by the equation:
5
ηεζµ =EOF (1.6)
where EOFµ is the electroosmotic mobility.
Surface of capillaryStern layer
Diffuse layer
Bulk solutionEOF
Surface of capillarySurface of capillaryStern layerStern layer
Diffuse layerDiffuse layer
Bulk solutionBulk solutionEOFEOF
Figure 1.2 Origin of EOF – Electrical Double Layer.
A unique characteristic of the EOF is that it is uniformly distributed along the capillary.
This results in a flat plug-like profile (Figure 1.3a) rather than a parabolic profile that is
generated by a pressure-driven system (Figure 1.3b), such as in high performance liquid
chromatography (HPLC). A flat profile of EOF reduces band broadening, enhances fast
elution rates, and yields highly efficient peaks.
Another distinctive feature of the EOF under normal conditions, where the
capillary surface is negatively charged, is that it causes movement of nearly all species in
the same direction, regardless of charge. In CZE, the migration of charged species is
6
determined by the EOF and by the electrophoretic mobility of the solutes. This can be
expressed by the following equation:
EOFepp µµµα += (1.7)
where ppαµ is the apparent mobility. The value of electrophoretic mobility of an ion
depends on the charge of that ion, and it can be either positive or negative. Therefore,
cations migrate fastest since both EOF and electrophoretic mobility are in the same
direction. Neutral molecules move along with the EOF, because they do not have
electrophoretic mobility, and they are not separated. Finally, anions migrate slowest since
their electrophoretic mobility is in the opposite direction of EOF. However, they still move
toward the detector, because the magnitude of EOF is much greater than their
electrophoretic mobility. This is clearly illustrated in Figure 1.4.
EOF
Flat profile
Pressure Driven Flow
Parabolic profile
Figure 1.3 (a) EOF flow profile, (b) Pressure driven flow profile and their corresponding solute zone profiles.
7
Capillary electrophoretic separations are also performed using another mode of CE,
called CGE. This form of CE is performed in a porous gel polymer matrix, and the
separation of analytes is based on their charge and size. CGE has been mostly employed
for the analysis of large biomolecules, such as proteins and nucleic acids. The most
common sieving gels used in CGE are cross-linked polyacrylamide and agarose. The
former gel has smaller pore size, and it is used for protein separations, while the latter, with
a larger pore size, is mostly applied to DNA analysis [2, 4, 5].
EOF
N
N
N
NN
N
EOF
N
N
N
NN
N
EOFEOF
NN
NN
NN
NNNN
NN
Figure 1.4 Solute migration.
CIEF is an electrophoretic technique used to separate amino acids, peptides, and
proteins based on their pI (isoelectric point) values. In CIEF, a pH gradient is formed
within the capillary using ampholytes. Ampholytes, or else zwitterions, are molecules that
contain both an acidic and a basic moiety. They have pI values between pH 3 and 9. The
8
gradient is formed when the capillary is filled with sample and ampholytes. A basic
solution is placed at the cathode, and an acidic solution is placed at the anode. When the
electric field is applied, the charged ampholytes and the sample components migrate
through the medium until they reach their pI value, at which they become uncharged. This
procedure is known as “focusing”, and it is indicated by a decrease in current. After this,
the contents of the capillary are mobilized to the detector by pressure or by adding an
electrolyte into one of the reservoirs [1, 4, 5].
CITP is another electrophoretic technique, in which a combination of two
electrolytes causes all analyte bands to migrate at the same velocity. This technique can
separate either cations or anions, but not both at the same time. During a separation
procedure, an analyte mixture is injected between a leading electrolyte containing ions of
higher mobility (Cl-) than any of the analyte ions and a terminating electrolyte with ions
that have lower mobility (heptanoate) than the sample ions. When an electric field is
applied, the analyte ions migrate in bands according to their unique mobilities. After all
analyte ions are separated into different bands, they move at the same velocity [1, 4].
MEKC and CEC are hybrid techniques that combine the best features of both
electrophoresis and chromatography. Both electrophoretic techniques can be used for the
separation of neutral as well as charged compounds. MEKC involves the introduction of a
surfactant at a concentration above the critical micellar concentration (CMC), at which
micelles are formed. The separation is based on the hydrophobic and ionic interactions of
the analytes with the micelles that act as a moving stationary phase, or else a
pseudostationary phase. CEC uses a stationary phase rather than a micellar
pseudostationary phase. As in MEKC, the mechanism of separation depends on the
9
partitioning between the two phases. However, in the case of CEC, the partitioning occurs
between the packed or coated stationary phase and the mobile phase. In addition, when the
analytes are charged, the separation also depends on their electrophoretic mobilities [3-7].
Both approaches are discussed in detail in the following sections.
1.2 Micellar Electrokinetic Chromatography
MEKC, which was first introduced by Terabe et al. in 1984 [8], is the second most
commonly used CE technique. Although MEKC is a form of CE, its separation principle is
more similar to HPLC than to CE. As stated earlier, the separation mechanism is based on
the electrophoretic mobility and the partitioning of the analytes between the mobile phase
and the micellar pseudostationary phase. However, for neutral species, it is only the
partitioning that effects the separation. The driving force for the partitioning of analytes is
hydrophobicity. In addition, hydrogen bonding, dipole-dipole, and dispersive interactions
can contribute to the solute partitioning between the two phases. The most commonly used
pseudostationary phase in MEKC is the micelle [1, 5, 9-11]. A more detailed discussion
about surfactants and micelles is given in the succeeding paragraphs.
Surfactants, which are also known as amphiphiles, are organic compounds that
consist of a polar or ionic head group and a hydrocarbon tail group (Figure 1.5). The head
group has hydrophilic properties, and it is compatible with the aqueous environment, while
the tail group is hydrophobic and not friendly with the aqueous environment. Surfactants
are classified into different groups according to the nature of their head group (anionic,
cationic, nonionic, and zwitterionic). Table 1.1 reports these four groups along with the
structures of different head groups and a few examples.
10
The surfactant molecules, at a low concentration, are dispersed in solution.
However, at a concentration above the CMC, and above a certain temperature, known as
the Krafft temperature, the surfactant molecules begin to build up their own structures.
They aggregate together to form micelles in the interior of the aqueous solution and
monolayers at the surface of the solution (Figure 1.6).
Hydrophobic Tail Group
Hydrophilic Head Group
Hydrophobic Tail Group
Hydrophilic Head Group
Figure 1.5 Structure of a surfactant molecule.
The CMC is a very important characteristic of each surfactant. Its value needs to be
determined in order to understand some of the characteristics and the properties of a
surfactant or a micelle. The CMC value may be attained graphically by plotting a
physicochemical property of the solution, i.e. surface tension, conductivity, turbidity,
osmotic pressure, fluorescence, and light scattering, versus the concentration of the
surfactant. The value of the CMC is the discontinuity in the plot, or the sudden variation in
the above relation. In an aqueous medium, above the CMC, surfactant molecules aggregate
together to form a micelle. As the surfactant concentration increases, the shape of the ionic
11
micelle changes from spherical to cylindrical, to hexagonal, and finally to lamellar. For
nonionic micelles, as the concentration increases the shape changes from spherical directly
to lamellar [6]. The number of aggregated molecules in a micelle is called the aggregation
number, n , and it is usually around 50 to 100. It can be determined by several analytical
techniques, such as light scattering, fluorescence, and NMR sedimentation.
Table 1.1 Classification of surfactant molecules used in MEKC.
ExamplesStructureHydrophilic group
SDSR-O-SO-3Anionic
CTAB, CTAC, DTABR-N+(CH3)3Cationic
R: alkyl groupFor abbreviations see pages xiv, xvi
PAPS, DAPSR-N+(CH3)2-(CH2)3-SO-3Zwitterionic
Triton X-100-RSR-Cyclohexyl-O-PEG
Octyl glucosideR-O-GlucoseNonionic
ExamplesStructureHydrophilic group
SDSR-O-SO-3Anionic
CTAB, CTAC, DTABR-N+(CH3)3Cationic
R: alkyl groupFor abbreviations see pages xiv, xvi
PAPS, DAPSR-N+(CH3)2-(CH2)3-SO-3Zwitterionic
Triton X-100-RSR-Cyclohexyl-O-PEG
Octyl glucosideR-O-GlucoseNonionic
The MEKC separation process for neutral analytes is only based on their
partitioning in and out of the micelle. When anionic micelles are used, the more the analyte
interacts with the micelle the longer its migration time. This is because anionic micelles
have electrophoretic mobilities that are in the opposite direction of the EOF. The more
12
hydrophobic analytes interact more strongly with the micelle, and they are retained longer
since the micelle partially offsets the effects of the EOF.
SurfactantMonolayer
Micelle
SurfactantMonolayer
Micelle
Figure 1.6 Equilibria between a surfactant molecule, a surface monolayer, and a micelle.
The separation mechanism in MEKC is chromatographic, and it can be described
by the use of modified chromatographic relationships. How effective a chromatographic
column is in separating two analytes partially depends on the relative rates at which the
analytes are eluted. These rates are determined by the magnitude of the equilibrium
constants (partition coefficients), K , which are defined as:
M
S
ccK = (1.8)
where Sc is the molar concentration of the analyte in the micellar pseudostationary phase
and Mc is its molar concentration in the mobile phase. The retention factor, or capacity
factor, 'k , describes the migration rates of analytes in the column, or the ratio of moles of
analyte in the micelle to those in the mobile phase:
13
( )
=
−
−=
M
S
m
r
r
VVK
ttt
ttk1
'
0
0 (1.9)
where rt is the retention time of the analyte, 0t is the retention time of the unretained
analyte that moves with EOF, mt is the retention time of the micelle, SV is the volume of
the micellar phase, and MV is the volume of the mobile phase. The above equation is partly
different from the normal chromatographic 'k because of the movement of the micellar
pseudostationary phase. If the micelle becomes the stationary phase the term mt becomes
infinite, and the Equation 1.9 is converted into the conventional chromatographic equation.
The selectivity factor α of a column for two analytes A and B is determined by the
equation:
( )( ) 0
0
tttt
Ar
Br
−−
=α (1.10)
where ( )Brt and ( )Art are the retention times of the more strongly and less strongly
retained analytes B and A, respectively. This definition dictates that the selectivity is
always greater than or equal to unity.
The resolution, SR , of a column describes its ability to separate two analytes. This
column resolution is defined as:
−
−
+
−
=
'1
0
0
'2
'2
2/1
1
1
11
4k
tttt
kkNR
m
mS α
α (1.11)
14
where N is the number of theoretical plates. This number of theoretical plates is widely
used as a quantitative measure of column efficiency, and is defined by use of the following
equation:
2
2/1
54.5
=
WtN r (1.12)
where 2/1W is the width of the peak at half its maximum height.
In Equation 1.11, the first derivative corresponds to efficiency, the second to
selectivity, and the third and fourth to retention. This suggests that resolution can be
improved by optimizing efficiency, selectivity, and/or the capacity factor. Since the
capacity factor is proportional to the concentration of the surfactant, it can be easily
modified by varying this concentration. In addition, resolution can be improved by
increasing the elution range, or time window, which is the time between 0t and mt (Figure
1.7). Neutral analytes elute between this elution range. The analytes that elute with the
EOF are hydrophilic and do not interact with the micelles. The analytes that are completely
retained by the micelles elute with the micelles. Maximum resolution is obtained when mt
t0
is very small (large time window). Selectivity can also be easily modified by varying the
size, charge, and geometry of the micelle, and particularly the surfactant [1, 3].
In chromatography and electrophoresis, resolution is given by:
( ) ( )[ ]BA
ArBrS WW
ttR
+−
=2
(1.13)
where AW and BW are the widths of the base of peaks A and B, respectively. A resolution
of 1.5 gives a baseline separation of the two analytes. The resolution can be improved by
15
increasing the length of the column, thus, increasing the number of theoretical plates
(column efficiency). However, this results in an increase in separation time [4].
0 t0 tr1 tr2 tr3 tm
Time
Time window
Analytes
EOF
Micelle
0 t0 tr1 tr2 tr3 tm
Time
Time window
Analytes
EOF
Micelle
Figure 1.7 Elution time window for neutral analytes in MEKC.
1.3 Capillary Electrochromatography
In recent years, CEC has become an important member of the arsenal of tools
available in separation science. Although the CEC approach was first introduced by
Pretorius et al. [12] in 1974, it received renewed interest during the 1990s [13-19]. This
revival of CEC is because it is a microcolumn electroseparation technique that combines
the best features of HPLC with those of CE. In essence, CEC mainly couples the high
16
selectivity of HPLC and the high separation efficiency of CE [20-26]. This is because in
CEC, the mobile phase solvent is transported through a capillary by use of the EOF. In
addition, CEC provides high resolution, short analysis time, ruggedness, and low sample
and mobile phase consumption. It also offers wider selectivity and facilitates the separation
of both neutral and charged compounds.
As discussed earlier, the latter characteristic is because the separation of analytes is
based on their interactions with the stationary phase and, when charged, their
electrophoretic mobility [15, 25-32]. Therefore, the apparent mobility for a charged analyte
in CEC is influenced by its electrophoretic mobility, its partitioning interaction and the
EOF. However, for a neutral analyte, the apparent mobility depends only on the
partitioning interaction with the stationary phase, while its elution is driven by the EOF.
Several manuscripts have described the capacity factor for separation of an analyte
in CEC [33-36]. Briefly, the capacity factor in CEC ( CECk ′ ) can be expressed as:
+
−
=
EOF
e
EOF
e
CEC
kk
µµµµ
1
'
' (1.14)
This equation incorporates both the electrophoretic and chromatographic separation
mechanisms in CEC. It should be noted that the mechanism of separation in CEC is highly
dependent upon the analyte. For neutral analytes, eµ is zero, and CECk ′ is equal to k ′ .
Thus, neutral analytes are solely separated on the basis of a chromatographic mechanism.
Finally, charged analytes are separated by a combination of chromatographic and
electrophoretic mechanisms, provided these analytes interact with the stationary phase.
17
In CEC, as in every chromatographic system, the capillary column is the most
important component. It serves as a vessel to transport the mobile phase and as a
separation channel [15, 28]. Therefore, preparation of the column, and particularly the
stationary phase packed or immobilized on the inner walls of the capillary, is critical for
CEC. The capillary columns in CEC are usually classified into three main formats [21, 22,
28, 37-44]: (i) packed-CEC [23-27, 45-52], (ii) open-tubular CEC (OT-CEC) [39-43, 53-
64], and (iii) monolithic (or continuous rod) CEC [20, 38, 65-85]. In this dissertation, the
first two main formats are discussed in detail.
1.3.1 Packed-Capillary Electrochromatography
In packed-CEC, a fused-silica capillary is filled with a typical HPLC packing
material (i.e. octadecyl silica). The mobile phase in packed-CEC is driven by the EOF that
is induced by applying an electric field across the capillary column. The origin of this flow
is the electrical double layer that is formed at the solid-liquid interface of a charged surface
in contact with the BGE (Figure 1.8).
In a capillary packed with silica particles the surfaces of the capillary wall and the
particles are negatively charged due to the dissociation of silanol groups. The velocity of
EOF in packed-CEC is shown in the following equation:
η
εεσµ
EcF
RTro
EOF
2/1
32
= (1.15)
where σ , which is proportional to zeta potential, is the charge density at the surface, oε is
the permittivity of vacuum (8.85 x 10-12 C2N-1µ-2), rε is the dielectric constant of the
mobile phase, R is the gas constant, T is temperature, c is the concentration of the
18
electrolyte, F is Faraday’s constant, η is the viscosity of the mobile phase, and E is the
electric field strength. The above equation illustrates that the velocity of EOF depends on
the charge density on the surfaces of the capillary walls and silica particles, the dielectric
constant, the viscosity, and the concentration of the BGE, and the temperature.
However, there are some problems that need to be solved in order for this
conventional form of CEC to be a viable alternative to both CE and HPLC. One of the
problems of packed-CEC is the requirement to fabricate frits, which are needed to retain
the packed particles inside the capillary column. Another problem in packed capillaries is
the tendency to form bubbles around the packing material or at the frit. This problem often
results in an unstable baseline, variable migration times, and current breakdown.
Pressurization of both ends of the column is required, and the mobile phase must be
thoroughly degassed in order to reduce the possibility of bubble formation. Another major
drawback of conventional CEC is that the packing procedure is generally more difficult
than for HPLC due to the narrow inner bore of the capillary and the small diameter of the
particles (5 µm or less). Finally, basic compounds are difficult to separate in packed-CEC
due to the presence of silanol groups that are needed to generate an adequate EOF [21, 39-
41, 86-93]. In order to circumvent the problems mentioned above, continuous bed-type
columns, i.e. open-tubular and monolithic columns, have been suggested as alternatives to
packed-CEC.
1.3.2 Open-Tubular Capillary Electrochromatography
In the OT-CEC format, the stationary phase is deposited on the inner walls of the
capillary. Preparation of the stationary phase, or coating, is a crucial step in any
chromatographic system. In OT-CEC, this coating needs to be stable in order to provide
19
efficient chromatographic separations and a reproducible EOF [92]. In OT-CEC, there are
six general approaches used for modifying the capillary column. These include (i)
adsorption, (ii) covalent bonding and/or crosslinking, (iii) porous silica layers, (iv)
chemical bonding after etching, (v) sol-gel, and (vi) molecular imprinting [39-42, 91-94].
Although OT-CEC is considered an alternative to packed-CEC, its phase ratio and sample
capacity are relatively low due to the typical small surface area of the coating. Several
options have been proposed to increase the surface area, and therefore, to increase the
interactions between the analyte and the coated phase. These options include polymer
coatings, porous silica layers, etching, and sol-gel techniques [22, 41, 89, 94].
Stationary phase particles
EOF
Electrical double layersCapillary wall
Stationary phase particles
EOF
Electrical double layersCapillary wall
Figure 1.8 Origin of EOF in a packed capillary.
20
1.3.2.1 Adsorption
Adsorption of chemicals to solid surfaces is a common phenomenon observed in
separation science. The adsorption of proteins, peptides, and basic compounds on the silica
capillary wall usually causes serious problems, such as loss in efficiency, peak tailing,
unstable baseline, nonreproducible migration times, and reduction of capillary lifetime. In
order to reduce or eliminate the analyte-wall interactions, different methods have been
developed. The most common methods shield the negatively charged silanol groups with
another layer, the adsorbed stationary phase [28, 39, 61, 95]. According to the strength of
adsorption, the adsorbed stationary phases can be divided into two groups: dynamically
adsorbed and physically adsorbed stationary phases [28, 30, 92]. If the adsorption of the
modifier on the capillary wall is weak, the stationary phase is called dynamically adsorbed
stationary phase. In this case, the modifier is added to the mobile phase. In contrast, if the
modifier is strongly adsorbed, the stationary phase is termed a physically adsorbed
stationary phase. In this case, the addition of the modifier to the mobile phase is not
necessary. Several modifiers have been used for the preparation of adsorbed stationary
phases [17, 18, 39, 59, 61, 62, 92, 93, 95-121]. These modifiers can be grouped into three
main categories: (i) cationic surfactants, (ii) polymeric surfactants, and (iii) charged
polymers.
(i) Cationic Surfactants
In 1990, Pfeffer and Yeung [17] developed a method for the separation of several
polycyclic aromatic hydrocarbons (PAHs) by use of open-tubular capillary liquid
chromatography (OT-CLC). The capillaries were first coated with a polymer solution of
0.9% PS-264 (polyvinylsiloxane). The velocity of the EOF on these columns was low due
21
to the polymer coating that blocked the silanol groups on the capillary wall. The cationic
surfactant cetyltrimethylammonium bromide (CTAB) was then added to the mobile phase
at low concentrations (µM). When this eluent was used in a polymer coated, reversed-
phase open-tubular column, a substantial EOF was developed. This, in turn, provided
highly efficient and fast separations of neutral compounds. In 1991, Pfeffer and Yeung
[18] used OT-CLC for the separation of anions. This separation involved partitioning of
the anions with the help of the ion-pairing agent, tetrabutylammonium (TBA) cation, being
adsorbed onto the surface of an open-tubular column. The separation of the anions was
based on differences in retention, rather than differences in electrophoretic mobility. The
ion pairing agent, which was added in the mobile phase at low concentrations (600 µM),
was used to control the affinity of the anions for the stationary phase.
Garner and Yeung [95] proposed the use of an ion-exchange mechanism for the
separation of compounds with similar mobilities, such as 4-amino-1-naphthalenesulphonic
acid and 5-amino-2-naphthalenesulphonic acid. Capillaries coated with a hydrophobic
stationary phase were shown to be dynamic ion exchangers when the quaternary
ammonium compound, CTAB, was added to the mobile phase. CTAB was dynamically
adsorbed to the column, forming a charged double layer. Sample ions were then retained
due to their coulombic attraction to the double layer. In this study, they showed that the
retention of the sample ions could be varied in this system by changing various parameters
such as the concentration of the buffer ion, the addition of an organic modifier, and the
concentration of the cationic surfactant CTAB.
Liu et al. [105-107] developed a novel preparation method for a physically
adsorbed stationary phase in OT-CEC for the separation of both achiral and chiral
22
compounds. The adsorbed stationary phases were prepared by simply rinsing the capillary
with a mobile phase containing a cationic surfactant, such as CTAB or a basic chiral
selector, such as protein, peptide or amino acid. After adsorption of the stationary phase,
the capillary was rinsed again with a mobile phase, which did not contain either CTAB or a
chiral selector. Five alkyl substituted benzenes were baseline separated within 6 min [105].
The run-to-run reproducibility of retention time was good with relative standard deviation
(RSD) values of less than 2.3%. However, the day-to-day reproducibility was not as good
with RSD values of less than 10% [106, 107].
Baryla et al. [100] described the adsorption mechanisms and aggregation properties
of the cationic surfactants CTAB and didodecyldimethylammonium bromide (DDAB) that
were used for the preparation of dynamically adsorbed stationary phases. Atomic force
microscopy (AFM) was used for the elucidation of the coating morphology. In addition,
separation of the basic proteins lysozyme, ribonuclease A, α-chymotrypsinogen A,
cytochrome c, and myoglobin was performed. Two of the five proteins were irreversibly
adsorbed to the wall. All three peaks that were present were broad and tailed due to wall
adsorption. The use of a CTAB-coated capillary gave a baseline separation of three of the
proteins in less than 15 min with high efficiencies. When a DDAB-coated capillary was
used all five proteins were separated in less than 6 min. This was due to the increased
surface coverage provided by DDAB.
(ii) Polymeric Surfactants
Although micelles have been successfully used in separations, they have some
significant limitations that cause problems in MEKC. The most important problem
associated with conventional micelles is the dynamic equilibrium between the surfactant
23
molecules and the micelle. The presence of dynamic equilibrium lowers the stability of the
pseudostationary phase. PAHs, which are considered extremely hydrophobic compounds,
are difficult to separate because they co-migrate with the micelle. This problem can be
solved by either increasing the concentration of the surfactant or by adding an organic
modifier in the BGE solution. However, organic modifiers usually perturb the micelle
structure. In addition, an increase in the concentration of the surfactant may cause longer
migration times and Joule heating, which has detrimental effects on MEKC separations [6,
7, 11]. In order to compensate the above problems, new pseudostationary phases have been
developed by use of chiral and achiral polymeric surfactants.
In 1994, Wang and Warner [120] reported the use of a polymeric surfactant added
to the BGE in MEKC. Polymeric surfactants offer several distinct advantages over
conventional micelles [121-126]. Firstly, polymerization of the surfactant eliminates the
dynamic equilibrium due to the formation of covalent bonds between the surfactant
aggregates. This, in turn, enhances stability and improves resolution. Secondly, polymeric
surfactants can be used at low concentrations because they do not depend on the CMC.
This usually provides higher efficiencies and rapid analysis.
Harrell et al. [119] developed a dynamic coating by use of a novel nonionic micelle
polymer, poly (n-undecyl-α-D-glucopyranoside). They applied this coating to the
separation of seven tricyclic antidepressants. Although physical and dynamic adsorptions
have simple and rapid coating procedures and good reproducibilities, they usually have
short lifetimes and limited pH ranges [92, 112, 113]. Both coatings are adsorbed to the
capillary wall via electrostatic interactions and hydrogen bonding. These interactions are
weaker than covalent bonds. However, multiple electrostatic interactions within a coating
24
that involves a layer-by-layer deposition process provides greater stability and longer
lifetime [92, 93, 112-114, 121].
A layer-by-layer coating is termed a polyelectrolyte multilayer (PEM), which is
usually constructed in situ by use of alternating rinses of positively and negatively charged
polymers [114, 127-130], where the negatively charged polymer may be a polymeric
surfactant [92, 93, 121]. A layer of polymer adds to the oppositely charged surface,
reversing the surface charge and priming the film for the addition of the next layer via
electrostatic forces. The advantages of such coatings are two-fold. First, less consumption
of the polymer is required, since it is adsorbed onto the capillary wall. Second, there is less
detection interference between the polymer and the analyte of interest, which in turn,
makes the system more amenable to coupling with mass spectrometry (MS).
Although the basic idea of the PEM coating is simple, a theoretical description is
relatively complex due to the long range of the coulombic interaction between layers. In
addition, several techniques have been employed (neutron reflectometry, atomic force
microscopy, infrared spectrometry) in order to better understand the mechanism of the
PEM coating formation. However, it is still not very well understood. The PEM coating is
constructed by the use of the simple procedure described above. The first layer should have
high adsorptivity in order to strongly attach the multilayer to the substrate. Several studies
have shown that after the first four or five layers, the thickness of the coating increases
linearly with the number of layers, and the zeta potential has a value of around zero. The
thickness of each layer depends on the amount of salt added to the polymeric solutions,
and not on the surface charge of the substrate. However, the total thickness of the coating
increases with a surface charge increase. In addition, it is observed that salt has the
25
strongest effect on the polyelectrolyte layer thickness. Polymer concentration, polymer
type, molar mass, salt type, deposition time, and solvent are less important variables [127,
131, 132].
All polymer deposition solutions contain an amount of salt, usually NaCl. Salt
moderates the electrostatic interactions that occur between the oppositely charged
polyelectrolyte sections, Pol+ and Pol-, according to the following chemical equation:
Pol+Pol-m + Na+
aq + Cl-aq ↔ Pol+Cl-
m + Pol-Na+m
where m refers to the multilayer phase. The charges on a polymer can be balanced by
either those on the oppositely charged polymer or by the salt counterions within the film.
In the first case, where the positive charge of one polymer is balanced by the negative
charge of another polymer, the compensation is called intrinsic (Figure 1.9a). In the second
case, the compensation is extrinsic, since much of the polymer charge is balanced by the
salt ions that are added in the polymer deposition solution (Figure 1.9b) [127, 131].
In this dissertation, the PEM coating approach is explored, and its performance is
evaluated by use of different drug compounds. For the achiral studies, the polymeric
surfactant poly (sodium N-undecanoyl-L-glycinate), poly (L-SUG), was used as the anionic
polymer and poly (diallyldimethylammonium chloride), PDADMAC, was used as the
cationic polymer [92]. A diagrammatic scheme of the PEM coated capillary is illustrated in
Figure 1.10. This diagram is not intended to give an actual structural representation of the
bilayer, but rather a representation of the order of polymer deposition. More details
concerning this work are given in Chapter 3.
26
+
-
- --
- --
----
--
-
+++
++
++++++
++
- --
- --
----
--
-
++
++ + + +
+ ++ + +
+
-
+
Na+
Na+
Na+
Na+
Cl- Cl- Cl-Cl-
(a) Intrinsic
(b) Extrinsic
+
-
- --
- --
----
--
-
+++
++
++++++
++
- --
- --
----
--
-
++
++ + + +
+ ++ + +
+
-
+
Na+
Na+
Na+
Na+
Cl- Cl- Cl-Cl-
(a) Intrinsic
(b) Extrinsic
Figure 1.9 (a) Intrinsic, (b) Extrinsic charge compensation.
Kamande et al. [93] investigated the use of a PEM coating for the separation of
phenols and benzodiazepines. For their studies, they used the polymeric surfactant, poly
(sodium undecylenic sulfate), poly (SUS), and PDADMAC in a single bilayer PEM
coating. The run-to-run and capillary-to-capillary reproducibilities were very good, and the
RSD values of EOF were less than 1.5%. The endurance of the coating was more than 100
runs. In addition, this study demonstrated the importance of the PEM coating by
comparing the separation of benzodiazepines using CZE and MEKC.
27
N+N+N+N+N+N+N+N+N+N+ N+N+N+N+N+N+N+N+N+N+N+ N+
CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NH
C O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-
Silicate surface Si
O-O(deprotonated
silanol groups)
Capillary wallSi
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-O
Si
O-
Silicate surface Si
O-O
Si
O-O(deprotonated
silanol groups)
Capillary wall
BilayerBilayer
Figure 1.10 Scheme of the PEM-coated capillary.
(iii) Charged Polymers
Erim et al. [104] reported the use of a simple method for the preparation of a
polyethyleneimine (PEI) coating on the inner surface of fused-silica capillaries. The PEI
layer was constructed by flushing the capillary with a solution that contained high-
molecular-mass PEI. These authors investigated the reproducibility and the long-term
stability of the PEI coating by measuring the plate numbers and retention times of some
basic proteins. The RSD values in migration times for run-to-run reproducibilities ranged
from 0.5 to 1.5%, and for column-to-column reproducibilities ranged from 1.9 to 2.8%.
The stability of the coating was tested by flushing the capillary with a solution at pH 11.0
for 60 hours. The migration times of all proteins after 60 hours slightly increased, and the
efficiencies were not altered.
A stable modification of the inner wall was also developed by using a simple
coating procedure, i.e. successive multiple ionic-polymer layer (SMIL) coating. Katayama
28
et al. [112, 113] used polybrene (PB) as the cationic polymer, and dextran sulfate (DS),
alginic acid, or hyaruronic acid as an anionic polymer. In their first study [112], they
established an anion-modified capillary (SMIL-DS capillary) by first attaching the cationic
polymer to the capillary wall, and then the anionic polymer to the cationic polymer layer.
Efficient separation of acidic proteins was achieved even at pH values under 7.4. The
ability to prevent the adsorption of the acidic proteins to the SMIL-DS capillary wall was
due to the presence of sulfonic groups in the DS layer. These groups provided strong
repulsion between the protein and the capillary wall. However, the SMIL-DS capillary was
not suitable for the separation of basic proteins since the capillary wall was negatively
charged through pH 2-11. In a pH range of 2-11, the SMIL-DS capillary exhibited a pH-
independent EOF from anode to cathode. The endurance of the SMIL-DS coated capillary
was more than 100 runs and the capillary-to-capillary variation was less than 1%. In their
next study [113], this group further modified the SMIL coating and developed a cation-
modified capillary (SMIL-PB capillary) by attaching the cationic polymer to the anionic
polymer layer. The SMIL-PB capillary was applied to the separation of basic proteins. The
separation was possible even when the pH of the mobile phase was near the pI of the
protein. The SMIL-PB capillary demonstrated strong endurance with the achievement of
600 runs and excellent chemical stability against 1 M sodium hydroxide (NaOH) and 0.1
M hydrochloric acid (HCl). The RSD values of the run-to-run, day-to-day, and capillary-
to-capillary reproducibilities were below 1%.
Poly (styrene sulfonate), PSS, is another anionic polymer that has been used in a
PEM coating procedure. Graul et al. [114] used this coating for the separation of a series of
basic proteins. The coatings used for protein separations and reproducibility studies
29
consisted of 6.5 layer pairs. A layer pair is a layer of cationic polymer plus a layer of
anionic polymer, also termed a bilayer. The polymer deposition solutions for the
construction of the first 3.5 bilayers contained no salt, and for the last 3 contained 0.5 M
sodium chloride (NaCl). They observed an excellent run-to-run stability over a wide range
of pH. These capillaries were also proven to be very stable to extremes of pH (pH 12.0 and
pH 2.0) and ionic strength, and to dehydration/rehydration. The RSD values of capillary-
to-capillary reproducibilities were less than 2% when both pH 6.0 and pH 4.0 were used.
In addition to the reproducibility control of EOF, these authors achieved stable flow rates
immediately after exposure of the column to mobile phase, as well as reverse flow.
1.3.2.2 Covalent Bonding and/or Crosslinking
Fixation of a layer by covalent bonding and/or crosslinking is another approach
used for modifying the capillary [58, 63, 101, 102, 133-142]. Although this approach
offers a long capillary lifetime, it usually requires a more complicated coating procedure.
Rehder et al. [58] studied the electrochromatographic separation of bovine β-lactoglobulin
variants A and B by covalently attaching DNA aptamer, which form a G-quartet
conformation to a capillary surface. Aptamers are short, single-stranded oligonucleotides
that are recognized for their high affinity and specific binding to target molecules. Their
use as a stationary phase offers many advantages. They are stable, easy and simple to
produce, manipulate, and attach to silica surfaces, and they are compatible with a wide
range of separation conditions and mobile phases. For the preparation of the stationary
phase, oligonucleotides were covalently attached to the inner capillary surface using the
organic linker molecule, sulphosuccinimidyl 4-(N-maleimidomethyl) cyclohexane-1-
carboxylate.
30
A porphyrin derivative, 5,10,15,20-tetrakis(penta fluorophenyl)porphyrin,
H2TPFPP, was also used as a capillary wall modifier in OT-CEC by Charvatova et al.
[102]. H2TPFPP was physically adsorbed or covalently bonded to the capillary surface for
the separation of aromatic carboxylic acids. In both covalently bonded and physically
adsorbed phases, the capillaries were filled with a solution of the porphyrin derivative in
dichloromethane. In the first case, both ends of the capillary were closed and the capillary
was left overnight at room temperature. In the second case, the capillary was left in the
vacuum oven for 2 hours at 60 °C. The synthetic strategy used for the preparation of
chemically bonded phases was based on the generation of anionic silanol groups on the
capillary wall, which were then employed for nucleophilic substitution at the para position
of H2TPFPP. The main advantage of H2TPFPP is that it is capable of increasing the EOF.
The results from this study showed that coating of the capillary with H2TPFPP gave a
better resolution at all pH values tested. Although both covalent bonding and physical
adsorption resulted in similar results at pH 8.5 and pH 6.0, covalently bonded capillaries
offered better results. At pH 5.0, physically adsorbed coatings lead to results, which were
not reproducible.
In another study, Chiari et al. [101] reported the use of poly (vinylamine), PVA, for
the separation of polyanionic acids. The coating procedure consisted of two steps: (i)
dynamic adsorption of PVA, and (ii) crosslinking and derivatization of PVA by
nucleophilic addition of primary amino groups to the conjugated double bonds of N, N-
methylenebisacrylamide and N, N, N-trimethylaminoethylacrylamide. The best stability of
this coating was achieved with the crosslinked PVA. There was no change in the EOF even
after 120 hours of continuous use at pH 4.0. RSD values and average transit times of ten
31
injections were taken every 6 hours during the 120-hour study. The values between 0.2 and
1% indicated excellent reproducibility.
Calixarene-coated capillaries were also used in OT-CEC. Such capillaries are
expected to provide more efficient separations of larger compounds [139-143]. Most
recently, Wu et al. [140] have successfully used p-tert-butylcalix[8]arene bonded
capillaries for the separation of o-, m-, and p-benzenediols, α- and β-naphthols, and α- and
β-naphthylamines. These capillaries were prepared with the use of γ-
glycidoxypropyltrimethoxysilane as a bridge. The bonded capillaries showed good
separation selectivity, suggesting significant interactions between the analytes and the
bonded phase. In addition, for the evaluation of the stability of the capillary surface, five
batches prepared over a period of six months were tested. All capillaries exhibited high
stability and reproducibility.
1.3.2.3 Porous Layers
Another approach to increasing the phase ratio, the surface area, and the loading
capacity of a capillary is by use of porous-layer open-tubular columns [13, 143-146].
Huang et al. [13] introduced the utility of such columns with a positively charged
hydrocarbonaceous porous layer as the stationary phase for the separation of basic proteins
and peptides. The porous layer was crosslinked and prepared by in situ polymerization of
vinylbenzyl chloride and divinylbenzene in the presence of 2-octanol as a porogen inside a
silanized fused-silica capillary. The chloromethyl groups at the surface of the porous layer
were reacted with N, N-dimethyldodecylamine to obtain a positively charged surface with
fixed C12 alkyl chains. Stability was monitored daily by measuring the electroosmotic
mobility. They observed that the migration times remained constant for more than a week.
32
A porous silica layer chemically modified with C18 groups was used as a stationary
phase by Crego et al. [143]. The 9.60 µm i.d. capillary used in this work had a thin porous
silica layer (0.70 µm) with C18 groups chemically bonded onto the prepared layer. These
authors investigated the effects of several experimental parameters, such as the volume
fraction and the type of organic modifier in the mobile phase, the concentration, type and
pH of the buffer on the electroosmotic mobility, the retention behavior of test analytes, and
the column efficiency. In their studies, they used a group of PAHs as a test mixture.
Consequently, they were able to optimize the separation of this group of analytes by
varying all of the operating parameters identified above.
The separation of charged analytes was also examined on porous-layer open-
tubular capillaries [146]. The preparation of the stationary phase was performed by use of
the sol-gel technique. This technique is discussed in detail later in this section. The
successful separation of acidic diuretics and N-alkylanilines as basic compounds at low pH
indicated that it is possible to separate charged compounds on porous-layer open-tubular
columns. Neutral aryl alkyl ketones were also successfully separated with high
efficiencies. However, basic pharmaceutical drugs demonstrated severe peak tailing and
were poorly separated by use of these columns. In addition, strongly acidic analytes were
not detected due to their high electrophoretic mobilities.
1.3.2.4 Chemical Bonding After Etching
Etching is another approach to overcoming the major problems associated with OT-
CEC. The etching process increases the surface area of the capillary by a factor of up to
1000 [147]. Therefore, more stationary phase can be attached to the wall, which in turn,
increases the loading capacity of the column. In addition, during the etching process,
33
dissolution and redeposition of silica material occurs. This creates radial extensions from
the wall that decrease the distance an analyte has to travel in order to interact with the
stationary phase [88]. The etching process was first introduced by Onuska et al. [147] for
gas chromatography (GC), and it was then modified for OT-CEC by Pesek et al. [14, 39,
41, 88-91, 148-156].
The entire procedure includes etching followed by chemical bonding. The etching
process uses ammonium hydrogen difluoride as the etching agent which, under controlled
conditions of temperatute and reaction time, can increase the surface area. The method of
bonding an organic moiety to the etched surface of a capillary utilizes the
silanization/hydrosilation method. In the first step, which involves silanization, the etched
surface is reacted with triethoxysilane (TES) to produce a hydride layer that is deposited on
the surface. In the second step, an organic moiety is attached to the hydride intermediate
via hydrosilation. This is accomplished by passing a solution that contains a terminal
alkene and a catalyst, typically hexachloroplatinic acid, through the capillary [92].
In a 1997 study, Pesek et al. [148] attached two organic moieties, octadecyl and
diol, to the etched capillary wall. Peptide (angiotensins) and protein samples were used for
comparison studies between these two columns and a bare capillary. The migration times
for each of the analytes increased in the following order: bare<diol<C18. Longer migration
times were observed when the C18-modified etched capillary was used. This was probably
due to stronger analyte-bonded phase interactions. From these studies, the authors
concluded that the bonded etched capillaries have significantly different retention
characteristics from a bare capillary. In addition, the separation capabilities of the two
34
modified capillaries varied due to differences in the chemical properties of the two organic
moieties.
Another study from Pesek et al. focused on a comparison of the migration behavior
of two antibiotics (ampicillin and gentamycin) between several types of etched open-
tubular capillaries [90]. The types of columns used included (i) etched and unmodified, (ii)
etched and bonded with an octadecyl moiety, (iii) etched and bonded with a cholesterol
moiety, (iv) etched and coated with anionic fluorosurfactant (FSA), and (v) etched and
coated with neutral zwitterionic fluorosurfactant (FSK). Each of the antibiotic samples
consisted of several components, and the resolution capabilities of the different types of
columns were compared. Their studies showed that each type of column gave a different
elution pattern due to the stationary phase effect. A number of variables had to be
optimized in order to get the maximum separation for these samples. Optimization was
achieved by varying several experimental conditions, such as the type of stationary phase,
the pH and composition of the buffer, and the applied voltage.
Matyska et al. [89] investigated the performance characteristics of n-octadecyl and
cholesterol- modified capillaries for the analyses of synthetic peptides. The results
indicated that the resolution and selectivity of peptides could be affected by varying the
type of chemically bonded group and the nature of the surface chemistry used to modify
the capillary wall. In addition, other experimental variables, such as the type and
percentage of organic solvent modifier, and the pH of the buffer system can affect the
retention of peptides. For the evaluation of the reproducibility and the stability of both
types of open-tubular columns, four batches were prepared over a period of two years. For
this study, a mixture of two small basic compounds, serotonin and tryptamine, and a
35
mixture of two basic proteins, chicken and turkey lysozyme, were used. The measurement
of the selectivities and efficiencies of these analytes showed that both the reproducibility
and stability were excellent. All of the etched modified capillaries used in this study still
performed well after more than 200 runs.
1.3.2.5 Sol-Gel Technique
As noted earlier, the preparation procedure for columns with expansive surface
areas and large loading capacities involves two steps: (i) etching or laying down a porous
layer, and (ii) attaching functional groups to the intermediate layer by chemical bonding
[30]. However, this two-step procedure may be time consuming. Another approach, which
involves the construction of polymeric stationary phases, involves good column stability
and high surface ratio. However, the main problem associated with these columns is the
poor column efficiency as a result of the slow diffusivity of analytes in the layers [145]. In
order to circumvent these drawbacks, the sol-gel technique was introduced for the
synthesis of organic-inorganic hybrid materials that are used as stationary phases in OT-
CEC. Recently, the sol-gel technique, which combines the synthesis of a bonded phase and
a supporting porous silica film in a single step, has gained much attention [29, 40, 145,
146, 157-164].
The advantages of the sol-gel processed stationary phases include high stability,
high mass loadability, great column efficiency, large surface area leading to higher
retentions, and a relatively simple preparation procedure [157, 159-161]. The sol-gel
process consists of five major steps: (i) hydrolysis, (ii) condensation and polycondensation
of sol-gel precursors, (iii) casting of the sol, (iv) aging, and (v) drying. In the first step, a
metal alkoxide is hydrolyzed under acidic or basic conditions to form the corresponding
36
metal hydroxide, which condenses and polycondenses to form “sol” particles. These “sol”
particles are then crosslinked to form a wet “gel,” which can take any desired shape. A dry
“gel,” or a “xerogel” is finally formed after the loss of solvent through aging and drying.
By controlling different processing parameters, such as temperature, pressure, alcohol
solvent, and pH during the different steps of the process, one can fabricate sol-gel derived
materials with specific structures [15, 157, 160-162].
In 1995, Guo and Colon [145] introduced the organosilicon sol-gel fabricated
coatings in OT-CEC. They synthesized a porous glass film in such a way that the
stationary phase (octyl groups) is incorporated into the glass matrix during the glass
formation process. This film was then used in capillaries, and functioned as a stationary
phase. The performance of this sol-gel derived stationary phase was evaluated by use of a
group of PAHs. Its stability was also studied under acidic and basic conditions. In order to
test the stability at low and high pH values, the capillary was washed with 1%
trifluoroacetic acid (pH ≈ 0.3) and with the mobile phase methanol/1 mM phosphate buffer
(70:30) at pH 11.4. Their results indicated that a retentive layer was still on the surface of
the capillary, even after exposure to high- and low-pH conditions. Finally, the mass
loadability of the column was evaluated by injecting different concentrations of
naphthalene in the capillary. The fact that the efficiency started to degrade at a
concentration of 100 mM indicated overloading. In a conventional column, the efficiency
deteriorated at concentrations below 10 mM.
Rodriguez and Colon [162] used n-octadecyltriethoxysilane (C18-TEOS) and
tetraethoxysilane (TEOS) as the precursors to fabricate an organic-inorganic hybrid
material by use of the sol-gel method. Fused-silica capillaries were coated with the C18-
37
TEOS/TEOS mixture and were tested using OT-CEC. A test mixture containing toluene,
ethylbenzene, biphenyl, dimethylnaphthalene and amybenzene was separated by using two
columns coated with (i) TEOS, and (ii) C18-TEOS/TEOS sol-gel derived materials.
Additionally, eight PAHs were separated by use of a C18-TEOS/TEOS-coated column.
These compounds were baseline separated only in the columns containing the organically
modified composite (C18-TEOS/TEOS).
A novel sol-gel approach for fabrication of open-tubular columns was described by
Hayes and Malik [158]. A surface-bonded octadecylsilane (ODS) stationary phase coating
was created by sol-gel chemistry. The use of a deactivating reagent, phenyldimethylsilane
(PheDMS), in the sol-gel solution was evaluated. The performance of both nondeactivated
and deactivated sol-gel columns was also evaluated with the test mixtures of PAHs,
benzene derivatives, and aromatic aldehydes and ketones. The column deactivation
resulted in a higher separation efficiency and better resolution. In addition, the RSD values
for the run-to-run reproducibility of the sol-gel columns were less than 0.70% and 0.80%
for the aromatic hydrocarbons and the aromatic carbonyl compounds, respectively.
Wang et al. [161] used 1,4,7,10-tetraazacyclotridecane-11,13, dione
(dioxo[13]aneN4) for the first time in the sol-gel approach for the preparation of
dioxo[13]aneN4 modified capillaries. In comparison with columns prepared by the sol-gel
process with just TEOS, the sol-gel derived macrocyclic dioxopolyamine columns were
able to give better separations of a mixture of isomeric nitrophenols and benzenediols, a
mixture of isomeric aminophenols and diaminobenzenes, and a mixture of four
neurotransmitters. The reproducibilities of migration times and plate numbers were
38
satisfactory with RSD values of less than 2% and less than 8.5% for the respective run-to-
run and column-to-column reproducibilities.
1.4 Chirality
Chirality is the geometric property that is responsible for the nonidentity of an
object with its mirror image [165]. In other words, an object is chiral if it is non-
superimposable on its mirror image. In 1848, Pasteur [166] marked the beginning of
chirality and chiral separation. He discovered that the resolution of the racemic mixture of
ammonium sodium tartrate yielded two enantiomorphic crystals. The separation was
perfomed by use of a pair of tweezers and a hand lens. He also observed that the crystals
gave a left and a right rotation of polarized light. In addition, he proposed that since the
difference of the optical rotation was examined in solution, the molecules are the ones that
are mirror images of each other [167].
Chiral compounds are molecules that relate to each other like a pair of hands. The
word chiral was derived from the Greek word cheir, which means hand. The chiral
molecules are also termed optically active because they have the ability to rotate plane-
polarized light. Optically active molecules that rotate light to the left are called
levorotatory (L), and they have the negative sign (-). If the molecules rotate the light to the
right, they are said to be dextrorotatory (D) or positive (+). In addition, chiral compounds
are also termed as molecules with non-superimposable mirror images. Molecules that are
mirror images of each other are also called enantiomers or optical isomers. These
molecules have an asymmetric tetrahedral carbon atom with four different substituents. If
the priority of the substituents is in a clockwise direction, the configuration is called R
(right or rectus). If the priority is in a counterclockwise direction, the configuration of the
39
chiral center is S (left or sinister). These differences are due to the asymmetric element in
the chiral molecule that can be a center, an axis, or a plane of asymmetry. An equimolar
mixture of two enantiomers is called a racemate, and it is designated by the symbol (±) [6,
165, 168].
Although enantiomers rotate plane-polarized light in opposite directions, these
molecules have identical physical properties such as boiling and melting points, and
spectroscopic properties such as nuclear magnetic resonance (NMR) spectra. However,
enantiomers in a racemic drug usually have different biological activities. For example,
dextromethorphan is an over-the-counter cough suppressant, whereas levomethorphan is a
controlled narcotic. Thalidomide is a sedative drug that was prescribed to pregnant women
in the early 1960’s. When this drug was taken during the early stages of pregnancy, it
prevented the normal growth of the fetus. This resulted in serious birth defects in
thousands of children around the world. It was later observed that the R-enantiomer was a
sedative, whereas the S form was the one that caused the foetal abnormalities. The
examples mentioned above illustrate the different pharmakokinetic characteristics and
pharmacological activities of each enantiomer in a racemic drug. Therefore, the
development of analytical techniques for the separation of chiral bioorganic molecules has
become very important [6, 168, 169].
In order to distinguish between two enantiomers in a racemic drug, a chiral selector
has to be added to the BGE solution. However, the mechanism of chiral discrimination is
not very well understood. In general, the “three point rule,” which was illustrated by
Easson and Stedman [170], describes the interactions that are necessary for chiral
discrimination. Chiral separation can be achieved if a minimum of three simultaneous
40
interactions occur between the chiral selector and one of the enantiomers. Due to spatial
restrictions, the other enantiomer should attain at least two of these interactions. These
interactions can be hydrophobic interactions between the hydrophobic core of the polymer
and the analyte, and electrostatic interactions between the polar head group of the polymer
and the analyte. Another type of interaction is the dipole-dipole forces, such as hydrogen
bonding between the polar group of the chiral selector and the analyte. In addition, other
secondary interactions can occur, such as π-π interactions, ion-dipole bonds, and Van der
Waals forces.
1.5 Chiral Selectors in Open-Tubular Capillary Electrochromatography
Chiral separation has received considerable attention in many industries,
particularly the pharmaceutical industry. Several analytical techniques have already been
developed for the separation of chiral bioorganic molecules. One of them is OT-CEC,
where the chiral selector is coated and immobilized onto the inner surface of a capillary. In
1992, Mayer and Schurig [171] reported the first enantiomeric separation in OT-CEC by
using capillaries coated with immobilized Chirasil-Dex (permethyl-β-cyclodextrin
chemically linked to dimethylpolysiloxane). In addition to derivatized cyclodextrins [164,
171-178], cellulose [55, 60], proteins [42, 105-107, 179-183], molecular imprinted
polymers (MIPs) [184-195], and polymeric surfactants [121] were also used.
1.5.1 Derivatized Cyclodextrins
Mayer et al. [171-174] used an immobilized Chirasil-Dex for the separation of a
number of chiral compounds. Chirasil-Dex is a permethyl-β- or γ-cyclodextrin covalently
bonded to a dimethylpolysiloxane via an octamethylene spacer. Chirasil-Dex modified
capillaries proved to be buffer-resistant and stable under neutral and acidic conditions.
41
Schurig et al. [175, 176] also investigated the use of an immobilized Chirasil-Dex
(mono-6-O-octamethylenepermethyl-β-cyclodextrin chemically linked to
dimethylpolysiloxane) in chromatography. The concept of unified enantioselective
chromatography was demonstrated for the chiral separation of hexobarbital by use of GC,
supercritical fluid chromatography (SFC), liquid chromatography (LC), and CEC [175].
The results from this study indicated that CEC is superior to GC, SFC and LC in regards to
several parameters, which are important in chiral separation. These parameters include the
chiral separation factor, α, peak resolution, Rs, and efficiency, N. In addition, Chiralsil-Dex
demonstrated a long lifetime, as well as configurational and thermal stability.
A related approach was also reported by Armstrong et al. [177] for the separation
of racemic mephobarbitol by OT-CEC. The capillary was coated with permethylated-β-
cyclodextrin that was covalently linked to dimethylpolysiloxane. This stationary phase
appeared to be stable and relatively unchanged by high temperature. Further, it could not
be removed from the capillary by conventional polar and nonpolar solvents or by
supercritical fluids.
Another approach for immobilization of β-cyclodextrin was developed by Pesek et
al. [178]. These authors etched the capillaries with ammonium hydrogendifluoride and
then modified them with a chiral selector by use of the silanization/hydrosilation method,
which was described earlier. The types of selectors evaluated include lactone, β-
cyclodextrin, and naphthylethylamine. For the etched cyclodextrin-modified capillary (2-
hydroxy-3-methacryloyloxypropyl-β-cyclodextrin), benzodiazepines were the only class of
compounds resolved among the test analytes used. They obtained partial resolution of the
two isomers for oxazepam, temazepam, chlorodiazepoxide, and diazepam. The stability of
42
the column was also good since it was used for at least 200 injections with no observed
changes in enantiomeric resolution.
Wang et al. [164] coated columns with 2,6-dibutyl-β-cyclodextrin (DB-β-CD)
using the sol-gel technique. This new technology provided columns that demonstrated
good resolution, stability, and reproducibility on separation of some positional isomers and
chiral compounds. The chiral compounds used in this study were ibuprofen and
binaphthol. The two isomers of both analytes gave partial resolution.
1.5.2 Cellulose
Capillaries coated with cellulose derivatives were demonstrated by Francotte et al.
[55] for the separation of a number of chiral pharmaceuticals. Their columns were prepared
by coating 3,5-dimethylphenylcarbamoyl cellulose (DMPCC) or para-methylbenzoyl
cellulose (PMBC) on 50 µm i.d. fused-silica capillaries. The capillaries were coated at 35-
40 °C and 0.3 mbar. A film thickness of about 0.025 µm was obtained by using a filtered
solution of the cellulose derivative in dichloromethane or tetrahydrofuran (THF) for
DMPCC or PMBC, respectively. Although the enantioselectivity of the coating was
excellent, the column stability was low, and its lifetime was relatively short. Unfortunately,
column lifetime rarely exceeded one hundred injections. These problems were largely due
to the fact that the cellulose coating was not immobilized on the inner wall of the capillary.
Recently, another study was performed by Wakita et al. [60], in order to examine
the enantioseparation ability of cellulose tris(3,5-dichlorophenylcarbamate), (CDCPC), in
OT-CEC. The capillary was filled with a solution containing CDCPC derivative, styrene,
dry THF, and a solution of 2,2-azobisisobutyronitrile. The column was then heated at 60
°C for 20 hours to produce copolymerization. The covalently bound CDCPC was used as a
43
chiral selector for the separations of trans-stilbeneoxide, laudanosine, etozolin, and
piprozolin. The enantiomers of the first two analytes were partially resolved and the
enantiomers of the last two analytes were baseline resolved on this CDCPC coated
capillary.
1.5.3 Proteins
Liu et al. [105-107] prepared chiral stationary phases for OT-CEC by use of a
method based upon the physical adsorption of a basic chiral selector such as protein,
peptide, and amino acid on the wall of a capillary. A number of chiral compounds were
successfully resolved using this system. The adsorbed protein, lysozyme, showed high
chiral selectivity, and the resolution ranged from 1.74 to 2.05.
Liu, Otsuka, and Terabe [42, 180-182] studied chiral separations by OT-CEC with
avidin as a chiral stationary phase. This coating was prepared by use of the physical
adsorption method that was proposed by Liu et al. [105-107]. A total of sixteen
enantiomeric compounds were separated with avidin adsorbed on the capillary wall [42,
180]. Among them, ten pairs of enantiomers were baseline separated, while the other
enantiomers were partly resolved with resolutions of 0.8-1.2. However, due to the low
phase ratio of OT-CEC, only enantiomers that have strong interactions with the stationary
phase can be separated. The RSD values for the run-to-run, day-to-day, and column-to-
column reproducibilities were less than 2.2%, 2.3%, and 1.1%, respectively. The column-
to-column reproducibility was better than the day-to-day reproducibility due to the small
loss of the stationary phase during the day-to-day reproducibility measurements. Therefore,
the capillary was rinsed daily with an avidin solution for 10 min prior to experiments.
44
The feasibility of OT-CEC for quantitative analysis of enantiomers was also
examined with an avidin adsorbed capillary [181]. With conventional injection, the lowest
detectable concentration was 10-6 M. The limits of detection (LODs) for enantiomers were
below 1%. The detection sensitivity was improved by use of the field-enhanced sample
injection (FESI) technique. In this technique, a water plug is introduced hydrodynamically
into the capillary inlet end, and then, the sample solution is introduced with electrokinetic
injection. Using the FESI technique, the LOD was reduced to a 10-9 M level, and the
enantiomeric ratio was improved to 0.3%. In addition to the sensitivity, both the efficiency
and resolution were improved.
Another approach to improving the sensitivity of OT-CEC is to use an extended
light path (ELP) capillary, which has a bubble cell at the detection point. A physically
adsorbed avidin stationary phase was used to evaluate an ELP capillary and an etched
capillary [182]. With an ELP capillary, the peak height was enhanced by a factor of as
much as 10. However, the peak efficiency and resolution decreased. When an etched
capillary was used, the phase ratio was slightly enhanced by a factor of 1.64, as compared
to an unetched capillary.
1.5.4 Molecular Imprinted Polymers
Over the last few years, molecular imprinting has attracted considerable attention as
an approach for the preparation of chiral stationary phases in OT-CEC. This is due to the
high enantioselectivity and predetermined elution order that characterizes the MIPs [44].
The selectivity of the resultant MIP is predetermined by the choice of template (imprint)
molecule used for the imprint preparation. Monomers are selected for their ability to
interact with the template molecule by mainly electrostatic interactions, hydrogen bonding,
45
dipole-dipole interactions, and hydrophobic interactions. The template molecule can either
be the analyte of interest or a structural analogue [194-196]. MIPs are prepared by
polymerization of a mixture that contains a template molecule, functional monomers, a
crosslinking agent, and an initiator in a nonpolar solvent (porogen). In the
prepolymerization mixture, the functional monomers interact with the imprint molecule,
and orient around it in a specific way to form complexes. During polymerization, in the
presence of a crosslinking agent, a rigid polymer is produced. After polymerization, the
imprint molecule can be removed by solvent extraction. This gives rise to a material that is
filled with cavities, and one that is complementary in size, shape, and chemical
functionality to the imprint molecule. These cavities enable the polymer to rebind the
original template molecule [189, 191, 193-197].
Schweitz and coworkers [189-192, 194, 195] prepared chiral stationary phases by
using molecular imprinting of the (R) and (S) enantiomers of propranolol. In 1997, Nilsson
et al. [189] performed the enantiomeric separation of β-adrenergic antagonists by using
three different capillary electrochromatographic methods. In these methods, they used
different cyclodextrins added to the mobile phase, a crosslinked protein gel, and a
molecularly imprinted ((R) enantiomer of propranolol) superporous polymer as chiral
selectors. All three methods were able to resolve or partially resolve all β-adenergic
antagonists into their enantiomers. Very rapid enantiomer separations of propranolol were
also studied by the use of short super-porous monolithic MIPs [194]. MIP stationary
phases were synthesized by use of an in situ photo-initiated polymerization reaction.
Propranolol enantiomers were resolved in less than 1 min. In addition, more MIP coatings
were synthesized by the use of a surface-coupled radical initiator [195]. A
46
prepolymerization mixture was prepared by dissolving the template molecule (S)
propranolol, the functional monomer methacrylic acid, and the crosslinking monomer
1,1,1-tris(hydroxymethyl)propane trimethacrylate in either toluene, dichloromethane, or
acetonitrile. The prepolymerization mixture was then introduced into the capillary and the
ends were sealed. The polymerization was performed by illuminating the capillary with a
UV lamp. The use of different solvents facilitates control of the coating, regarding its
morphology and appearance. Furthermore, all MIP coatings synthesized using the different
solvents, provided enantiomeric separation of propranolol when the (S) enantiomer was
used as the template molecule.
Tan et al. [194] reported a method for in situ preparation of molecular imprint
polymers as thin films inside 25 µm i.d. fused-silica capillaries. Methacrylic acid and 2-
vinyl pyridine were used as the functional monomers. Ethylene dimethacrylate or
trimethylol propane trimethacrylate was used as the crosslinker and toluene as the porogen.
Chiral separations of the enantiomers D- and L- dansyl phenylalanines were achieved in
both OT-LC and OT-CEC. The resolution in OT-CEC was much higher than in OT-LC.
1.5.5 Polymeric Surfactants
In this dissertation, the chiral separations of 1,1’-binaphthyl-2,2’-
dihydrogenphosphate (BNP), 1,1’-bi-2-naphthol (BOH), secobarbital, pentobarbital and
temazepam are investigated by using the PEM coating [121]. However, the PEM coating
procedure used in the achiral studies [92] needed to be modified in order to achieve chiral
separations. PDADMAC was used as the cationic polymer, and the polymeric surfactant
poly (sodium N-undecanoyl-L-leucylvalinate), poly (L-SULV), was used as the anionic
polymer. Thus far, poly (L-SULV) has shown the best chiral discrimination ability for a
47
number of pharmaceutical compounds [126]. The PEM coating approach applied to chiral
separations is discussed in detail in Chapter 4.
1.6 Laser Scanning Confocal Microscopy
Since the beginning of observational science, the need to observe objects smaller
than those visible by eye alone has driven the development of microscopy. Conventional or
“far-field” light microscopy is the oldest form of microscopy. Very early in the
development of microscopy, it was realized that the spatial resolution that could be
achieved was limited by diffraction phenomena. This limit is best known as the diffraction
limit, or Abbe’s limit. Abbe showed that the theoretical resolution limit for an objective
lens is approximately half the wavelength, λ , of light employed. This relationship is
demonstrated in the following equation:
NAR λ61.0= (1.16)
where NA is the numerical aperture of the objective lens. The numerical aperture of an
objective lens is a measure of the angular size of the focusing cone of light and is defined
as:
θsinnNA = (1.17)
where θ is the acceptance angle of the lens, and n is the refractive index of the medium
(usually air, n=1.0) between the sample and objective lens. Therefore, the resolution in far-
field optical microscopes depends on the wavelength of light used. The use of a shorter
wavelength light can improve the spatial resolution. However, when ultraviolet light is
used several difficulties arise. These low wavelengths are usually absorbed by atmospheric
48
gases, liquids and many of the materials from which conventional optical elements are
constructed [198, 199].
In order to eliminate the problems caused by the far-field optics, and to improve the
optical resolution, near-field scanning optical microscopy (NSOM) was developed. Synge,
in 1928, was the first to propose the development of NSOM as a scanned probe
microscopy. He proposed the use of a light source formed from a subwavelength-size
aperture in a metal screen (Figure 1.11). When light passes through the backside of the
aperture, fields are produced in and near the front side of the hole. Only the sample region
directly beneath the aperture is illuminated. Images can be formed by moving the aperture
and sample relative to one another (raster scanning). The signal is observed by the use of a
single-element detector. The resolution is determined by the size of the aperture and its
distance from the sample, rather than the wavelength of the light [198].
Confocal microscopy is a valuable tool for obtaining three-dimensional images of
thick objects. It is widely used in the fluorescence mode for imaging biological species and
in the reflection mode for imaging objects of different forms. In the fluorescence mode, the
objects are stained with fluorescent dyes. Fluorescent dyes are molecules that absorb light
at one wavelength and emit light at another wavelength. When these molecules absorb
light at a specific absorption wavelength, the electron rises to a higher energy level, called
the excited state. In this state, electrons are unstable; thus, they return to the ground state
by releasing energy in the form of heat and light. This emission of energy is called
fluorescence. Due to some loss in energy in the form of heat, the emitted light has less
energy. Therefore, the light is emitted at a longer wavelength than the absorbed light [200,
201].
49
Near Field
Metal ScreenAperture
<< λλλλ
Incident Plane Waves
Sample
<< λλλλ
Far Field λ
X
Y
Z
Near Field
Metal ScreenAperture
<< λλλλ
Incident Plane Waves
Sample
<< λλλλ
Far Field λ
X
Y
Z
Y
Z
Figure 1.11 Model for the aperture-based near-field optical microscope.
Figure 1.12 is a schematic diagram of a confocal microscope. This microscope
differs from a conventional optical microscope in that it images points of light rather than
large volumes of light. In general, it illuminates and images the object one point at a time
through a pinhole. Light from a laser source passes through a small pinhole, which acts as
a spatial filter, to an objective lens. This lens focuses the light onto a small spot on the
object, at the focal plane of the objective lens. The light that is reflected back from the
illuminated object is collected by the objective lens, and it is partially reflected by a beam
splitter. Then, it is directed at a confocal aperture, or pinhole, that is placed in front of the
detector (i.e. photomultiplier tube). The pinhole rejects light that did not originate from the
50
focal plane of the microscope objective lens. Therefore, only a very small volume of light
is focused at any time. This is what gives the system its confocal property [200, 202].
The laser scanning confocal microscope (LSCM) is the most widely used confocal
microscope in the areas of biomedicine and material science. It works in both transmission
and reflection modes. In addition, it gives higher resolution and thinner optical sections
than those obtained by use of the conventional microscope. By scanning a lot of thin
sections through the sample, a very clear three-dimensional image of the specimen can be
produced. Scanning is important in LSCM because only a small volume is illuminated at a
time, but a larger area should be collected for producing a better image. Scanning can be
accomplished by either beam scanning or stage scanning. The advantage of beam scanning
is that a total image can be generated from the small volumes of light without moving the
specimen. During beam scanning the specimen is stationary, and it is scanned by flying
spots of light. However, the advantage of stage scanning over beam scanning is that the
total light intensity is much greater.
The most important components of a typical LSCM system are a light source, an
objective lens, a scanning device, pinhole lens, a beam splitter, a detector, an analog signal
processor, a digital signal processor, an image analyzer, and a computer. In LSCM, the
light source should be a stable source of spatially coherent light. Therefore, the single
mode laser is most commonly used for this purpose. For fluorescence applications, multi-
line lasers, such as Ar, KrAr, and HeNe, can be used. The objective lens is used both to
illuminate and receive the reflected light from the illuminated spot on the specimen. After
the laser beam is converged by the objective lens and focused on the specimen, the scanner
scans the focused spot in an x, y raster pattern in order to produce an image of the sample
51
point by point. The reflected light from the specimen is collected by the objective lens, and
reflected off by the beam splitter to the pinhole and the detector. The pinhole is an
important component for the determination of both the axial ( 2
2)(NA
nzR λ= ) and lateral
(NA
yxR λ4.0),( = ) resolution of the microscope. When the pinhole is large, it transmits
more light to the detector. This, in turn, generates a larger signal with lower resolution.
However, a smaller pinhole results in a higher resolution and a lower signal-to-noise ratio.
After the light passes through the pinhole, it is directed to the detector, which generates an
electrical signal. Then, the electrical signal is amplified and sent to a scan converter, where
it is combined with the x and y position signals in order to produce an image for display
[203-205].
1.7 Scope of Dissertation
In this dissertation, both packed and open-tubular capillary electrochromatographic
methods were developed for the achiral and chiral separations of various classes of
analytes. Chapter 2 is a report of studies on the packed mode of capillary
electrochromatography (CEC). The goal of this work was to separate seven
benzodiazepines by the use of a 40 cm packed bed of Reliasil 3 µm C18 stationary phase.
Optimal conditions were established by varying the mobile phase, the amount of organic
modifier, the buffer concentration, the applied voltage, and the column temperature. In
addition, the method developed was applied to the characterization of oxazepam in a
standard urine sample.
52
Beam Splitter
Objective Lens
ObjectFocal Plane
Light Source
Source Pinhole
Detector Pinhole
Photodetector
In-focus light
Out-of-focus light
Figure 1.12 Schematic diagram of a confocal microscope.
In Chapter 3, the open-tubular mode of CEC, which is an alternative approach to
conventional CEC, is described. In this approach, a polyelectrolyte multilayer (PEM)
coating procedure was used to construct thin films on the capillary walls of fused silica
capillaries. For the fabrication of this coating both positively and negatively charged
polymers were utilized. Poly (diallyldimethylammonium chloride), PDADMAC, was used
as the cationic polymer and poly (sodium N-undecanoyl-L-glycinate), poly (L-SUG), was
53
used as the anionic polymer. These capillaries were evaluated by use of seven
benzodiazepines as analytes. The run-to-run, day-to-day, week-to-week and capillary-to-
capillary reproducibilities were evaluated by computing the relative standard deviations
(RSDs) of electroosmotic flow (EOF). In addition, the chromatographic performance of the
monomeric form of the polymeric surfactant was compared for the separation of these
analytes.
Chapter 4 is an outline of an investigation of the chiral separations of 1,1’-
binaphthyl-2,2’-dihydrogenphosphate (BNP), 1,1’-bi-2-naphthol (BOH), secobarbital,
pentobarbital and temazepam by using the PEM coating approach and the polymeric
surfactant poly (sodium N-undecanoyl-L-leucylvalinate), poly (L-SULV), as the chiral
discriminator. However, the PEM coating procedure used in the achiral studies (Chapter 3)
needed to be modified in order to achieve chiral separations. The quaternary ammonium
salt PDADMAC was used as the cationic polymer, and the polymeric surfactant poly (L-
SULV) was used as the anionic polymer. Optimal conditions were established by varying
the additive (sodium chloride, 1-ethyl-3-methyl-1H-imidazolium hexafluorophosphate, 1-
butyl-3-methylimidazolium tetrafluoroborate) in the polymer deposition solutions, the salt
concentration, the column temperature, and the bilayer number. Reproducibilities were
also evaluated by using the RSD values of the EOF and the first peak (R-(+)-BNP).
Chapter 5 details an approach that utilizes open-tubular capillary
electrochromatography (OT-CEC) and laser scanning confocal microscopy (LSCM) to
further investigate the stability of the PEM coating after exposure to 0.1 M and 1.0 M
NaOH. The multilayer coatings used for these studies consisted of two and twenty layer
pairs. The structural changes of these coatings were monitored and imaged using LSCM
54
after flushing the capillaries with 1.0 M NaOH. This technique also allowed a study of the
uniformity and discontinuity of the coating. Using OT-CEC, both electroosmotic mobility
and selectivity changes were measured after flushing the capillaries with 0.1 M and 1.0 M
NaOH. In addition, a correlation between flushing time and change in electroosmotic
mobility and selectivity was examined.
1.8 References
1. Heiger, D. N. In High Performance Capillary Electrophoresis – An Introduction, 2nd ed.; Hupe, P., Rozing, G., McManigill, D., van de Goor, T., Swedberg, S., Jegle, U., Verhoef, M., Jarke, L., Ford, J., Krull, B., Eds.; Hewlett-Packard Company: France, 1992.
2. Terabe, S.; Otsuka, K.; Nishi, H. J. Chromatogr. A 1994, 666, 295. 3. Chiari, M.; Nesi, M.; Righetti, P. G. In Capillary Electrophoresis in Analytical
Biotechnology; Righetti, P. G., Ed.; CRC Press, Inc.: Boca Raton, FL, 1996. 4. Skoog, D. A.; Holler, J. F.; Nieman, T. A. In Principles of Instrumental Analysis,
5th ed.; Sherman, M., Bortel, J., Messina, F., Eds.; Harcourt Brace College Publishers: Orlando, FL, 1998.
5. Chankvetadze, B. In Capillary Electrophoresis in Chiral Analysis; John Wiley & Sons Ltd: London, UK, 1997.
6. Williams, C. C.; Shamsi, S. A.; Warner, I. M. In Advances in Chromatography;
Brown, P. R., Grushka, E., Eds.; Marcel Dekker, Inc.: New York, NY, 1997, 363-423.
7. Yarabe, H. H.; Billiot, E.; Warner, I. M. J. Chromatogr. A 2000, 875, 179. 8. Terabe, S.; Otsuka, K.; Ichikawa, K.; Tsuchiya, A.; Ando, T. Anal. Chem. 1984,
56, 111. 9. Terabe, S.; Otsuka, K.; Ando, T. Anal. Chem. 1985, 57, 834.
10. Terabe, S.; Ozaki, K.; Otsuka, K.; Ando, T. J. Chromatogr. 1985, 332, 211. 11. Haynes, J. L.; Shamsi, S. A.;Warner, I. M. Anal. Chem. 1999, 18, 317.
12. Pretorius, V.; Hopkins, B. J., Schieke, J. D. J. Chromatogr. 1974, 99, 23.
55
13. Huang, X.; Zhang, J.; Horvath, C. J. Chromatogr. A 1999, 858, 91. 14. Pesek, J. J.; Matyska, M. T. J. Chromatogr. A 1996, 736, 255. 15. Malik, A. Electrophoresis 2002, 23, 3973. 16. Knox, J. H.; Grant, I. H. Chromatographia 1991, 32, 317. 17. Pfeffer, W. D.; Yeung, E. E. Anal. Chem. 1990, 62, 2178. 18. Pfeffer, W. D.; Yeung, E. S. J. Chromatogr. 1991, 557, 125. 19. Hindocha, D.; Smith, N. W. J. Chromatogr. A 2000, 904, 99. 20. Chen, Z.; Hobo, T. Electrophoresis 2001, 22, 3339. 21. Liu, Z.; Otsuka, K.; Terabe, S.; Motokawa, M.; Tanaka, N. Electrophoresis 2002,
23, 2973. 22. Tsuda, T.; Matsuki, S.; Munesue, M.; Kitagawa, S., Oehi, H. J. Liq Chromatogr.
& Rel. Techn. 2003, 26, 697. 23. Chen, X.; Zou, H.; Ye, M.; Zhang, Z. Electrophoresis 2002, 23, 1246. 24. Deng, Y.; Zhang, J.; Tsuda, T.; Yu, P. H.; Boulton, A. A.; Cassidy, R. M. Anal.
Chem. 1998, 70, 1586. 25. Henry, C. W.; McCarroll, M. E.; Warner, I. M. J. Chromatogr. A 2001, 905, 319. 26. Thiam, S.; Shamsi, S. A.; Henry, C. W.; Robinson, J. W.; Warner, I. M. Anal.
Chem. 2000, 72, 2541. 27. Colon, L. A.; Guo, Y.; Fermier, A. Anal. Chem. 1997, 69, 461A. 28. Liu, Z.; Wu, R.; Zou, H. Electrophoresis 2002, 23, 3954. 29. Wu, J. T.; Huang, P.; Li, M. X.; Qian, M. G.; Lubman, D. M. Anal. Chem. 1997,
69, 320. 30. Zou, H.; Ye, M. Electrophoresis 2000, 21, 4073. 31. Lammehofer, M.; Lindner, W. J. Chromatogr. A 1999, 839, 167. 32. Lammehofer, M.; Lindner, W. J. Chromatogr. A 1998, 829, 115.
56
33. Wu, J. T.; Huang, P.; Li, M. X.; Lubman, D. M. Anal. Chem. 1997, 69, 2908. 34. Ye, M.; Zou, H.; Liu, Z.; Ni, J.; Zhang, Y. Science in China B 1999, 42, 639. 35. Walhagen, K.; Unger, K. K.; Olsson, A. M.; Hearn, M. T. W. J. Chromatogr. A
1999, 853, 263. 36. Zou, H.; Ye, M. Electrophoresis 2000, 21, 4073. 37. Paces, M.; Kosek, J.; Marek, M.; Tallarek, U.; Seidel-Morgenstern, A.
Electrophoresis 2003, 24, 380. 38. Wistuba, D.; Schurig, V. Electrophoresis 2000, 21, 3152. 39. Pesek, J. J.; Matyska, M. T.; Sentellas, S.; Galceran, M. T.; Chirai, M.; Pirri, G.
Electrophoresis 2002, 23, 2982. 40. Constantin, S.; Freitag, R. J. Chromatogr. A 2000, 887, 253. 41. Matyska, M. T.; Pesek, J. J.; Katrekar, A. Anal. Chem. 1999, 71, 5508. 42. Liu, Z.; Otsuka, K.; Terabe, S. J. Sep. Sci. 2001, 24, 17. 43. Jinno, K.; Sawada, H.; Trend in Anal. Chem. 2000, 19, 664. 44. Wistuba, D.; Schurig, V. J. Chromatogr. A 2000, 875, 255. 45. Tobler, E.; Lammerhofer, M.; Lindner, W. J. Chromatogr. A 2000, 875, 341. 46. Ye, M.; Zou, H.; Lei, Z.; Wu, R.; Liu, Z.; Ni, J. Electrophoresis 2001, 22, 518. 47. Tegeler, T.; Rassi, Z. E. Electrophoresis 2002, 23, 1217. 48. Huber, C. G.; Choudhary, G.; Horvath, C. Anal. Chem. 1997, 69, 4429. 49. Bailey, C. G.; Yan, C. Anal. Chem. 1998, 70, 3275. 50. Pirogov, A. V.; Buchberger, W. J. Chromatogr. A 2001, 916, 51. 51. Lim, L. T.; Zare, R. N.; Bailey, C. G.; Rakestraw, D. J.; Yan, C. Electrophoresis
2000, 21, 737. 52. Wen, E.; Asiaie, R.; Horvath, Cs. J. Chromatogr. A 1999, 855, 349.
53. Charvatova, J.; Deyl, Z.; Kasicka, V.; Kral, V. J. Chromatogr. A 2003, 990, 159.
57
54. Fujimoto, C. Electrophoresis 2002, 23, 2929. 55. Francotte, E.; Jung, M. Chromatographia 1996, 42, 521. 56. Charvatova, J.; Kral, V.; Kasicka, V. Collection Symposium Series 2001, 4, 109. 57. Pesek, J. J.; Matyska, M. T. J. Capillary Electrophoresis 1997, 4, 213. 58. Rehder, M. A.; McGown, L. B. Electrophoresis 2001, 22, 3759. 59. Tan, Z. J.; Remcho, V. T. Anal. Chem. 1997, 69, 581. 60. Wakita, T.; Chankvetadze, B.;Yamamoto, C.; Okamoto, Y. J. Sep. Sci. 2002, 25,
167. 61. Charvatova, J.; Matejka, P.; Kral, V.; Deyl, Z. J. Chromatogr. A 2001, 921, 99. 62. Sun, P.; Landman, A.; Barker, G. E.; Hartwick, R. A. J. Chromatogr. A 1994,
685, 303. 63. Mingxian, H.; Guoliang, Y.; Bradshow, J. S.; Milton, L. J. Microcol. Sep. 1993,
5, 199. 64. Guan, N.; Zeng, Z.; Wang, Y.; Fu, E.; Cheng, J. Anal. Chim. Acta 2000, 418, 145. 65. Kato, M.; Dulay, M. T.; Bennett, B.; Chen, J. R.; Zare, R. N. Electrophoresis
2000, 21, 3145. 66. Fujimoto, C.; Fujise, Y. Anal. Chem. 1996, 68, 2753. 67. Kornysova, O.; Owens, P. K.; Maruska, A. Electrophoresis 2001, 22, 3335. 68. Koide, T.; Ueno, K. J. Chromatogr. A 2000, 893, 177.
69. Gusev, I.; Huang, X.; Horvath, C. J. Chromatogr. A 1999, 855, 273. 70. Kang, J.; Wistuba, D.; Schurig, V. Electrophoresis 2002, 23, 1116. 71. Vegvari, A.; Foldesi, A.; Hetenyi, C.; Kocnegarova, O.; Schmid, M. G.;
Kudirkaite, V.; Hjerten, S. Electrophoresis 2000, 21, 3116. 72. Schmid, M. G.; Grobuschek, N.; Tuscher, C.; Gubitz, G.; Vegvari, A.;
Machtejevas, E.; Maruska, A.; Hjerten, S. Electrophoresis 2000, 21, 3141.
73. Chen, Z.; Hobo, T. Anal. Chem. 2001, 73, 3348.
58
74. Lammerhofer, M.; Peters, E. C.; Yu, C.; Svec, F.; Frechet, J. M. J.; Lindner, W. Anal. Chem. 2000, 72, 4614.
75. Lammerhofer, M.; Svec, F.; Frechet, J. M. J.; Lindner, W. Anal. Chem. 2000, 72,
4623. 76. Koide, T.; Ueno, K. Anal. Sci. 2000, 16, 1065. 77. Koide, T.; Ueno, K. J. Chromatogr. A 2001, 909, 305. 78. Peters, E. C.; Petro, M.; Svec, F.; Frechet, J. M. J. Anal. Chem. 1997, 69, 3646. 79. Peters, E. C.; Petro, M.; Svec, F.; Frechet, J. M. J. Anal. Chem. 1998, 70, 2288. 80. Peters, E. C.; Petro, M.; Svec, F.; Frechet, J. M. J. Anal. Chem. 1998, 70, 2296. 81. Que, A. H.; Novothy, M. V. Anal. Chem. 2002, 74, 5184. 82. Wu, R.; Zou, H.; Ye, M.; Lei, Z.; Ni, J. Electrophoresis 2001, 22, 544. 83. Wu, R.; Zou, H.; Fu, H.; Jin, W.; Ye, M. Electrophoresis 2002, 23, 1239. 84. Ping, G.; Schmitt-Kopplin, P.; Hertkorn, N.; Zhang, W.; Zhang, Y.; Kettrup, A.
Electrophoresis 2003, 24, 958. 85. Legido-Quigley, C.; Marlin, N. D.; Melin, V.; Manz, A.; Smith, N. W.
Electrophoresis 2003, 24, 917. 86. Baltussen, E.; van Dedem, G. W. K. Electrophoresis 2002, 23, 1224. 87. Smith, N. W.; Carter-Finch, A. S. J. Chromatogr. A 2000, 892, 219. 88. Pesek, J. J.; Matyska, M. T. J. Chromatogr. A 2000, 887, 31.
89. Matyska, M. T.; Pesek, J. J.; Boysen, R. I.; Heam, M. T. W. Anal. Chem. 2001,
73, 5116. 90. Pesek, J. J.; Matyska, M. T.; Tran, H. J. Sep. Sci. 2001, 24, 729.
91. Jinno, K.; Sawada, H.; Catabay, A. P.; Watanabe, H.; Sabli, N. B. H.; Pesek, J. J.;
Matyska, M. T. J. Chromatogr. A 2000, 887, 479.
92. Kapnissi, C. P.; Akbay, C.; Schlenoff, J. B.; Warner, I. M. Anal. Chem. 2002, 74, 2328.
59
93. Kamande, M. W.; Kapnissi, C. P.; Zhu, X.; Akbay, C.; Warner, I. M. Electrophoresis 2003, 24, 945.
94. Xie, M. J.; Feng, Y. Q.; Da, S. L.; Meng, D. Y.; Ren, L. W. Anal. Chim. Acta
2001, 428, 255. 95. Garner, T. W.; Yeung, E. S. J. Chromatogr. 1993, 640, 397. 96. Breadmore, M. C.; Macka, M.; Avdalovic, N.; Haddad, P. R. Anal. Chem. 2001,
73, 820. 97. Breadmore, M. C.; Palmer, A. S.; Curran, M.; Macka, M.; Avdalovic, N.;
Haddad, P. R. Anal. Chem. 2002, 74, 2112. 98. Breadmore, M. C.; Macka, M.; Haddad, P. R.; Avdalovic, N. Analyst 2000, 125,
1235. 99. Boyce, M. C.; Breadmore, M.; Macka, M.; Doble, P.; Haddad, P. R.
Electrophoresis 2000, 21, 3073. 100. Baryla, N. E.; Melanson, J. E.; McDermott, M. T.; Lucy, C. A. Anal. Chem.
2001, 73, 4558.
101. Chiari, M.; Ceriotti, L.; Crini, G.; Morcellet, M. J. Chromatogr. A 1999, 836, 81.
102. Charvatova, J.; Kasicka, V.; Deyl, Z.; Kral, V. J. Chromatogr. A 2003, 990, 111.
103. Breadmore, M. C.; Boyce, M.; Macka, M.; Avdalovic, N.; Haddad, P. R. J. Chromatogr. A 2000, 892, 303.
104. Erim, F. B.; Cifuentes, A.; Poppe, H.; Kraak, J. C. J. Chromatogr. A 1995, 708,
356.
105. Liu, Z.; Zou, H.; Ni, J. Y.; Zhang, Y. Anal. Chim. Acta 1999, 378, 73.
106. Liu, Z.; Zou, H.; Ye, M.; Ni, J.; Zhang, Y. Chinese J. Chromatogr. 1999, 17, 245.
107. Liu, Z.; Zou, H.; Ye, M.; Ni, J.; Zhang, Y. Electrophoresis 1999, 20, 2891.
108. Cifuentes, A.; Poppe, H.; Kraak, J. C.; Erime, F. B. J. Chromatogr. B 1996, 681, 21.
109. Gilges, M.; Kleemiss, M. H.; Schomburg, G. Anal. Chem. 1994, 66, 2038.
110. Cordova, E.; Gao, J.; Whitesides, G. M. Anal. Chem. 1997, 69, 1370.
60
111. Chiu, R. W.; Jimenez, J. C.; Monnig, C. A. Anal. Chim. Acta 1995, 307, 193.
112. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 2254.
113. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 5272.
114. Graul, T. W.; Schlenoff, J. B. Anal. Chem. 1999, 71, 4007.
115. Sawada, H.; Jinno, K. Electrophoresis 1999, 20, 24.
116. Bendahl, L.; Hansen, S. H.; Gammelgaard, B. Electrophoresis 2001, 22, 2565.
117. Kohr, J.; Engelhardt, H. J. Microcol. Sep. 1991, 3, 491.
118. Li, M. X.; Liu, L.; Wu, J. T.; Lubman, D. M. Anal. Chem. 1997, 69, 2451.
119. Harrell, C. W.; Dey, J.; Shamsi, S. A.; Foley, J. P.; Warner, I. M. Electrophoresis 1998, 19, 712.
120. Wang, J.; Warner, I. M. Anal. Chem. 1994, 66, 3773.
121. Kapnissi, C. P.; Valle, B. C.; Warner, I. M. Anal. Chem. 2003, 75, 6097.
122. Yarabe, H. H.; Billiot, E.; Warner, I. M. J. Chromatogr. A 2000, 875, 179.
123. Palmer, C. P.; Tanaka, N. J. Chromatogr. A 1997, 792, 105.
124. Palmer, C. P.; Terabe, S. Anal. Chem. 1997, 69, 1852.
125. Wang, J.; Warner, I. M. J. Chromatogr. A 1995, 711, 297.
126. Shamsi, S. A.; Valle, B. C.; Billiot, F.; Warner, I. M. Anal. Chem. 2003, 75, 379.
127. Schlenoff, J. B.; Ly, H.; Li, M. J. Am. Chem. Soc. 1998, 120, 7626.
128. Schlenoff, J. B.; Dubas, S. T.; Farhat, T. Langmuir 2000, 16, 9968.
129. Laurent, D.; Schlenoff, J. B. Langmuir 1997, 13, 1552.
130. Rmaile, H. H.; Schlenoff, J. B. Langmuir 2002, 18, 8263.
131. Dubas, S. T.; Schlenoff, J. B. Macromolecules 1999, 32, 8153.
132. Castelnovo, M.; Joanny, J.-F. Langmuir 2000, 16, 7524.
61
133. Huang, X.; Horvath, C. J. Chromatogr. A 1997, 788, 155.
134. Cifuentes, A.; de Frutos, M.; Santos, J. M.; Diez-Masa, J. C. J. Chromatogr. 1993, 655, 63.
135. Schmalzing, D.; Piggee, C. A.; Foret, F.; Carrilho, E.; Karger, B. L. J.
Chromatogr. 1993, 652, 149.
136. Towns, J. K.; Regnier, F. E. J. Chromatogr. 1990, 516, 69.
137. Bao, J. J. J. Liq. Chromatogr. & Rel. Tecnol. 2000, 23, 61.
138. Towns, J. K.; Bao, J. M.; Regnier, F. E. J. Chromatogr. 1992, 599, 227.
139. Zeng, Z.; Xie, C.; Li, H.; Han, H.; Chen, Y. Electrophoresis 2002, 23, 1272.
140. Wu, X.; Liu, H.; Liu, H.; Zhang, S.; Haddad, P. R. Anal. Chim. Acta 2003, 478, 191.
141. Li, H. B.; Zeng, Z. R.; Xie, C. H.; Chen, Y. Y. Chromatographia 2002, 55, 591.
142. Wang, Z.; Chen, Y.; Yuan, H.; Huang, Z.; Liu, G. Electrophoresis 2000, 21,
1620.
143. Crego, A. L.; Martinez, J.; Marina, M. L. J. Chromatogr. A 2000, 869, 329.
144. Bruin, G. J. M.; Tock, P. P. H.; Kraak, J. C.; Poppe, H. J. Chromatogr. 1990, 517, 557.
145. Guo, Y.; Colon, L. A. Anal. Chem. 1995, 67, 2511.
146. Dube, S.; Smith, R. M. Chromatographia 2003, 57, 485.
147. Onuska, F. I.; Comba, M. E.; Bistricki, T.; Wilkinson, R. J. J. Chromatogr. 1977,
142, 117.
148. Pesek, J. J.; Matyska, M. T.; Mauskar, L. J. Chromatogr. A 1997, 763, 307.
149. Catabay, A. P.; Sawada, H.; Jinno, K.; Pesek, J. J.; Matyska, M. T. J. Cap. Elec. 1998, 005:1/2, 89.
150. Pesek, J. J.; Matyska, M. T.; Cho, S. J. Chromatogr. A 1999, 845, 237.
151. Pesek, J. J.; Matyska, M. T.; Swedberg, S.; Udivar, S. Electrophoresis 1999, 20,
2343.
62
152. Matyska, M. T.; Pesek, J. J.; Yang, L. J. Chromatogr. A 2000, 887, 497.
153. Matyska, M. T.; Pesek, J. J.; Sandoval, J. E.; Parkar, U.; Liu, X. J. Liq. Chrom. & Rel. Technol. 2000, 23, 97.
154. Matyska, M. T.; Pesek, J. J.; Boysen, R. I.; Hearn, M. T. W. J. Chromatogr. A
2001, 924, 211.
155. Matyska, M. T.; Pesek, J. J.; Boysen, I.; Hearn, T. W. Electrophoresis 2001, 22, 2620.
156. Pesek, J. J.; Matyska, M. T. J. Chromatogr. A 1996, 736, 313.
157. Narang, P.; Colon, L. A. J. Chromatogr. A 1997, 773, 65.
158. Hayes, J. D.; Malik, A. Anal. Chem. 2001, 73, 987.
159. Zhao, Y.; Zhao, R.; Shangguan, D.; Liu, G. Electrophoresis 2002, 23, 2990.
160. Rodriguez, S. A.; Colon, L. A. Anal. Chim. Acta 1999, 397, 207.
161. Wang, Y. C.; Zeng, Z. R.; Xie, C. H.; Guan, N.; Fu, E. Q.; Cheng, J. K.
Chromatographia 2001, 54, 475.
162. Rodriguez, S. A.; Colon, L. A. Applied Spectroscopy 2001, 55, 472.
163. Crosnier de Bellaistre, M.; Mathieu, O.; Randon, J.; Rocca, J. L. J. Chromatogr. A 2002, 971, 199.
164. Wang, Y.; Zeng, Z.; Guan, N.; Cheng, J. Electrophoresis 2001, 22, 2167.
165. Jacques, J.; Collet, A.; Wilen, S. H. In Enantiomers, Racemates, and Resolutions;
Krieger Publishing Co: Malabar, FL, 1981.
166. Pasteur, L. Ann. Chim. Phys. 1848, 24, 442.
167. Feibush, B.; Grinberg, N. In Chromatographic Chiral Separations – The History of Enantiomeric Resolution, 2nd ed.; Zief, M., Crane, L. J., Eds.; Marcel Dekker, Inc.: New York, NY, 1988.
168. Ahuja, S. In Chiral Separations – Applications and Technology; American
Chemical Society: Washington, DC, 1997.
169. http://chirality.ouvaton.org/homepage.htm
63
170. Easson, E. H.; Stedman, E. J. Biochem. 1933, 27, 1257.
171. Mayer, S.; Schurig, V. J. High Res. Chromatogr. 1992, 15, 129.
172. Mayer, S.; Schurig, V. J. Liq. Chromatogr. 1993, 16, 915.
173. Mayer, S.; Schurig, V. Electrophoresis 1994, 15, 835.
174. Mayer, S.; Schleimer, M.; Schurig, V. J. Microcol. Sep. 1994, 6, 43.
175. Schurig, V.; Jung, M.; Mayer, S.; Fluck, M.; Negura, S.; Jakubetz, H. J. Chromatogr. A 1995, 694, 119.
176. Jakubetz, H.; Czesla, H.; Schurig, V. J. Microcol. Sep. 1997, 9, 421.
177. Armstrong, D. W.; Tang, Y.; Ward, T.; Nichols, M. Anal. Chem. 1993, 65, 1114.
178. Pesek, J. J.; Matyska, M. T.; Menezes, S. J. Chromatogr. A 1999, 853, 151.
179. Yang, J.; Hage, D. S. Anal. Chem. 1994, 66, 2719.
180. Liu, Z.; Otsuka, K.; Terabe, S. Chromatogr. 2000, 21, 302.
181. Liu, Z.; Otsuka, K.; Terabe, S. Electrophoresis 2001, 22, 3791.
182. Liu, Z.; Otsuka, K.; Terabe, S. J. Chromatogr. A 2002, 961, 285.
183. Hofstetter, H.; Hofstetter, O.; Schurig, V. J. Microcol. Sep. 1998, 10, 287.
184. Lin, J. M.; Uchiyama, K.; Hobo, T. Chromatographia 1998, 47, 625.
185. Quaglia, M.; de Lorenzi, E.; Sulitzky, C.; Massolini, G.; Sellergren, B. Analyst
2001, 126, 1495.
186. Bruggemann, O; Freitag, R.; Whitcombe, M. J.; Vulfson, E. N. J. Chromatogr. A 1997, 781, 43.
187. Lin, J. M; Nakagama, T.; Uchiyama, K.; Hobo, T. J. Liq. Chromatogr. & Rel.
Techn. 1997, 20, 1489.
188. Lin, J. M.; Nakagama, T.; Uchiyama, K.; Hobo, T. J. Pharmac. Biomed. Analysis 1997, 15, 1351.
189. Nilsson, S.; Schweitz, L.; Peterson, M. Electrophoresis 1997, 18, 884.
64
190. Schweitz, L.; Spegel, P.; Nilsson, S. Analyst 2000, 125, 1899.
191. Schweitz, L.; Andersson, L. I.; Nilsson, S. Anal. Chem. 1997, 69, 1179.
192. Spegel, P.; Schweitz, L.; Nilsson, S. Electrophoresis 2001, 22, 3833.
193. Tan, Z. J.; Remcho, V. T. Electrophoresis 1998, 19, 2055.
194. Schweitz, L.; Andersson, L. I.; Nilsson, S. Anal. Chim. Acta 2001, 435, 43.
195. Schweitz, L. Anal. Chem. 2002, 74, 1192.
196. de Boer, T.; Mol, R.; de Zeeuw, R. A.; de Jong, G. J.; Sherrington, D. C.; Cormack, P. A. G.; Ensing, K. Electrophoresis 2002, 23, 1296.
197. Quaglia, M.; de Lorenzi, E.; Sulitzky, C.; Caccialanza, G.; Sellergren, B.
Electrophoresis 2003, 24, 952.
198. Higgins, D. A.; Mei, E. In Near-Field Scanning Optical Microscopy; Bonnell, D., Eds.; Wiley-VCH, Inc: New York, NY, 2001, 371-420.
199. http://www.jasco.co.uk/nearfield.htm
200. Sheppard, C. J. R. In Multidimensional Microscopy; Cheng, P. C., Lin, T. H.,
Wu, W. L., Wu, J. L., Eds.; Springer-Verlag New York Inc.: New York, NY, 1994, 1-31.
201. http://dept.kent.edu/projects/cell/FLUORO.HTM
202. Rochow, T. G.; Tucker, P. A. In Introduction to Microscopy by Means of Light,
Electrons, X rays, or Acoustics, 2nd ed.; Plenum Press: New York, NY, 1994.
203. Sheppard, C. J. R. In Multidimensional Microscopy; Cheng, P. C., Lin, T. H., Wu, W. L., Wu, J. L., Eds.; Springer-Verlag New York Inc.: New York, NY, 1994, 53-71.
204. Corle, T. R.; Kino, G. S. In Confocal Scanning Optical Microscopy and Related
Imaging Systems; Academic Press: San Diego, CA, 1996.
205. http://www.cs.ubc.ca/spider/ladic/intro.html
65
CHAPTER 2.
SEPARATION OF BENZODIAZEPINES BY USE OF CAPILLARY ELECTROCHROMATOGRAPHY
2.1 Introduction
Benzodiazepines are effective medicinal compounds, which are used primarily for
the treatment of anxiety and sleep disorders [1]. In fact, these drugs are among the most
widely prescribed of all psychoactive drugs. They are important in forensic toxicology
having hypnotic, tranquillizing and anticonvulsant properties. Therefore, they are often
encountered in casework involving road traffic offences or drug overdose [2-4]. They enter
the brain rapidly and work by binding to a specific type of receptor protein, which is
widely distributed in groups of nerve cells involved in anxiety, memory, sedation, and
coordination. In recent years, there has been a growing interest regarding the side effects of
benzodiazepines. These effects include dizziness, adverse interaction with alcohol, and risk
of dependence after long-term use [5]. For these reasons, the analysis of such compounds
is vital to areas such as pharmaceutical analysis, therapeutic drug monitoring, and forensic
toxicology [6-8]. A variety of methods have been developed for the separation of
benzodiazepines, but only some of them have been applied to their identification and
determination in complex matrices such as blood [9-21], urine [22-27] and hair [28, 29].
The methods most frequently used for controlling and monitoring these drugs in
blood or urine are voltametry [30, 31], radioimmunoassay [6, 15], spectrophotometry [32],
supercritical fluid chromatography (SFC) [33], thin layer chromatography (TLC) [25, 34],
gas chromatography (GC) [10, 11, 26-28] and high-performance liquid chromatography
(HPLC) [2, 16-21, 34-38]. Among these techniques, GC and HPLC have been the most
66
popular due to their selectivity. However, GC analysis is complicated and time consuming,
due to the need for derivatization and the thermal instability of some drugs such as
oxazepam [39]. According to the literature, the separation of benzodiazepines is mainly
performed by liquid chromatography (LC) using reversed-phase systems composed of
silica support materials and chemically bonded alkyl chains (octyl or octadecyl) [2, 34]. In
comparison with existing electrokinetic techniques such as capillary zone electrophoresis
(CZE) and micellar electrokinetic chromatography (MEKC), HPLC has low column
efficiency due to the need for pressure-driven flow.
CZE and MEKC are well established for the separation of many classes of
compounds. However, CZE is unable to resolve benzodiazepines, because the majority of
them are neutral and are of similar hydrophobicity. MEKC has been proposed as an
alternative approach. In this approach, micelles are introduced into the background
electrolyte and the separation of some neutral species is achieved [40, 41]. MEKC has been
successfully used to separate benzodiazepines [13, 22-24, 39, 42]. Boonkerd et al. [42]
have studied the migration behavior of a series of benzodiazepines in MEKC using three
kinds of surfactants, i.e., sodium dodecyl sulfate (SDS), dodecyl trimethylammonium
bromide (DoTAB) and bile salts. Renou-Gonnord and David [39] have studied the effects
of SDS, β-cyclodextrin (β-CD), urea, organic solvents and applied voltage on the
separation of nine benzodiazepines using MEKC. Although easy to implement, MEKC
lacks the selectivity and the variety of stationary phases that HPLC offers [43, 44].
Another disadvantage of MEKC is its incompatibility with mass spectrometry detection
due to the high concentrations of surfactants used [45].
67
CEC is considered as an alternative approach to MEKC. It uses a stationary phase
rather than a micellar pseudo-stationary phase. Solutes are separated according to their
partitioning between the mobile and stationary phase and, when charged, their
electrophoretic mobility [46, 47]. The mobile phase in CEC is driven by electroosmotic
flow (EOF) induced by applying an electrical field over the column [45]. The advantages
that CEC offers over HPLC include higher efficiency, the possibility of using small
particles in beds that would create high back pressure in HPLC, and the unique selectivity
due to the superimposition of chromatographic and electrophoretic effects [48].
CEC has been successfully used for the analysis of neutral drugs [49-51]. However,
only a few studies have been performed on the separation of benzodiazepines using CEC
[44, 45, 52, 53]. Cahours et al. [45] have investigated the influence of temperature, ionic
strength and organic modifier content on electrophoretic, chromatographic and separation
performances of five benzodiazepines using CEC on a phenyl-bonded silica column. Jinno
et al. [52] have compared the separation behavior of a series of benzodiazepines in packed
CEC and open-tubular CEC using a cholesteryl silica-bonded phase.
In this chapter, the influence of several experimental parameters is described in
order to obtain improved selectivity and efficiency for the separation of seven
benzodiazepine standards. This is accomplished by use of an octadecyl silica (ODS)
stationary phase in CEC. The optimized method proved to be effective in characterizing
oxazepam in a urine sample.
68
2.2 Experimental
2.2.1 Reagents and Chemicals
Oxazepam, lorazepam, nitrazepam, clonazepam, temazepam, flunitrazepam,
diazepam and tris(hydroxymethyl)aminomethane·hydrochloride (Tris·HCl) were purchased
from Sigma Chemical Company (St Louis, MO, USA). Acetonitrile (ACN), methanol
(MeOH) and tetrahydrofuran (THF) were obtained from Fisher (Springfield, NJ, USA).
Hydrochloric acid was purchased from Mallinckrodt & Baker (Paris, KY, USA).
Polyimide-coated fused-silica capillary columns of 100 µm i.d. × 365 µm o.d. were
obtained from Polymicro Technologies (Phoenix, AZ, USA). The columns were packed
with 3 µm Reliasil CEC C18 stationary phase that was purchased from Column Engineering
(Ontario, CA, USA). The DAT Multi-Drug High Urine Calibrator was donated by Earl K.
Long hospital (Baton Rouge, LA, USA).
2.2.2 Instrumentation and Conditions
The CEC experiments reported here were conducted using an HP3DCE capillary
electrophoresis system (Agilent Technologies, Wilmington, DE, USA). Data were
collected by use of HP3DCE Chemstation software (Agilent Technologies). The dimensions
of the capillary were 48 cm × 100 µm i.d. (40 cm to the detector). All CEC separations
were performed by pressurizing both the inlet and outlet buffer vials with 12 bar of
nitrogen to prevent bubble formation. Unless stated otherwise, the temperature of the
capillary cassette was maintained at 25 °C by the instrument thermostating system.
Detection of the analytes was performed using a photodiode array detection system set to
220 nm. All sample solutions were injected electrokinetically (15 kV for 5 s).
69
2.2.3 Sample and Buffer Preparation
Analytical standard benzodiazepine stock solutions were prepared at concentrations
of 2 mg/ml in MeOH. A 400 µl aliquot of each analyte was mixed and the final
concentration of each benzodiazepine in the test mixture was ∼ 0.3 mg/ml. A buffer
solution of 100 mM Tris was prepared by dissolving the appropriate amount of Tris buffer
in 10 ml of deionized water and the pH was adjusted to 8.0 using 1M HCl. An appropriate
percentage of ACN, MeOH or THF (v/v) was added to an appropriate percentage of the
aqueous Tris buffer solution (v/v), and then, the final volume was adjusted with deionized
water depending on the mobile phase being studied. The final solution was filtered using a
polypropylene nylon filter with 0.45 µm pore size and sonicated for 15 min. Finally, the
urine sample was injected into the CEC capillary without any preparation.
2.2.4 Preparation and Conditioning of Packed Capillary Columns
The CEC columns were packed in our laboratory according to a standard procedure
developed elsewhere [54, 55]. The 3 µm Reliasil CEC C18 silica stationary phase was
slurried in acetone at a concentration of 0.2 g/ml. After sonication, 1 ml of slurry was
injected through a Rheodyne injector connected to a stainless steel reservoir. The injector
was connected to the pump that was a Knauer pneumatic HPLC pump (Berlin, Germany)
and the slurry reservoir was connected to the capillary. The other end of the capillary was
connected to a union containing a 0.5 µm frit. The pump pressure was set to 400 bar. When
the capillary was filled with stationary phase, the pump was turned off and the excess
slurry was removed from the reservoir. The capillary was reconnected and the pump was
set back to 400 bar for two more hours. While the pump was on, the first frit was
fabricated using an electrically heated Nichrome wire. The bottom union from the capillary
70
was removed and the excess stationary phase was flushed out with 200 bar pump pressure.
The last procedure was repeated after the preparation of the second frit. The detection
window was placed adjacent to the outlet frit by burning off the polyimide coating. The
packed capillary column was flushed with the mobile phase for one hour and then installed
in a capillary cartridge. The column was further conditioned by applying both pressure of
12 bar to the inlet side and potential in 5 kV increments for 10 min up to 25 kV. Finally,
both the inlet and the outlet vials were pressurized, and the voltage was set to 30 kV until
the current was stabilized. This procedure was used whenever a new mobile phase was
tested. Between injections, the CEC capillary columns were conditioned for 5 min using 10
kV.
2.3 Results and Discussion
The chemical structures and the numerical designations for each of the seven
benzodiazepines used in this study are shown in Figure 2.1. The influence of operating
parameters, such as nature and amount of organic modifier, buffer electrolyte
concentration, applied voltage, and temperature was studied to optimize the CEC
separation of these benzodiazepines.
2.3.1 Effect of Organic Modifier
Mobile phases comprised of Tris (10 mM, pH 8) modified with 60%, 70% and 45%
of either ACN, MeOH or THF, respectively, were used to study the influence of the
modifier on the separations of benzodiazepines. The organic solvent content was fixed in
order to have mobile phases with the same elution strength. A nomograph, which provides
the interconversion of reversed-phase mobile phases having the same strength, can be
found elsewhere [56, 57]. Vertical lines in this figure intersect mobile phases having the
71
same strength. For example, 70% MeOH has the same strength as 60% ACN or 45% of
THF. The buffer Tris was chosen because its low mobility would more closely match that
of the analytes, when compared to more conventional buffers such as borate and
phosphate. The low mobility of Tris also allows higher concentrations of buffer to be used
without significantly increasing the current. In addition, the relatively high ionic strength
of the buffer leads to sharper and more defined peaks. The ionic strength of each mobile
phase in this study was constant (10 mM). A pH of 8 was chosen because the EOF at this
pH had a greater stability and the analysis time was shorter as compared to lower pH
values.
N
HN
OH
O
Cl N
HN
OH
O
Cl
Cl
N
HN
O2N
O
N
HN
O
O2N
Cl
N
N
OH
O
Cl
H3C
N
NO
O2N
F
H3C
N
N
Cl
OH3C
1. Oxazepam 2. Lorazepam 3. Nitrazepam 4. Clonazepam
5. Temazepam 6. Flunitrazepam 7. Diazepam
Figure 2.1 Structures of the seven benzodiazepine analytes.
72
For a given capillary, the EOF ( EOFµ ) is defined as:
0VtLL td
EOF =µ (2.1)
where dL is the distance from injector to detector, tL is the total capillary length, 0t is the
migration time of the electroosmotic flow marker and V is the applied voltage. The
relative EOF can be monitored by use of the values of 0t , since all the other factors are
constant as is evident from Equation 2.1. In the studies reported here, the values of 0t were
measured with MeOH.
Figure 2.2 demonstrates electrochromatograms for the separation of
benzodiazepines on a C18 stationary phase using Tris-ACN (40:60), Tris-MeOH (30:70)
and Tris-THF (55:45) binary mixtures. A reduction in electroosmotic mobility was
observed from 1.78×10-4 for the ACN-buffer mixture to 4.64×10-5 cm2V-1s-1 for THF–
buffer mixture. When ACN-buffer mixture was used, the total separation time decreased
and the peak efficiencies increased at the expense of lower resolution and retention factor
values. Using Tris-THF (55:45), the total separation was very long and the peak
efficiencies were low. Although the analytes in the mixture were not well resolved using
Tris-ACN (40:60), they eluted in 20 min. Taking into consideration the peak efficiency and
the speed of analysis, the Tris-ACN (40:60) binary mixture was used to further optimize
separation conditions for the benzodiazepines.
The retention mechanism of benzodiazepines on a C18 stationary phase with a
mobile phase of ACN/H2O is based on the differential partitioning of the analytes into the
alkyl-bonded phase. Their retention is determined by hydrophobic interactions between the
C18 stationary phase and the nonpolar moiety of each analyte, and by interactions between
73
the polar mobile phase and the sample molecules. The migration order of benzodiazepines
for the ACN/H2O mobile phase was tr1,2< tr
3,4< tr5< tr
6< tr7. However, the elution order
changed when MeOH and THF were used. The elution order for the MeOH/H2O mobile
phase is tr3,4< tr
2< tr1< tr
6< tr5< tr
7 and tr1,2< tr
5< tr6< tr
3< tr4< tr
7 for the THF/H2O mobile
phase.
2.3.2 Effect of Mobile-Phase Composition
In an attempt to achieve baseline separation, the content of the aqueous buffer (10
mM Tris, pH 8) was increased from 30% to 60%. As depicted in Figure 2.3, both
selectivity and retention time of the analytes increase with decreasing ACN concentration.
A decrease from 70% to 40% ACN increased the selectivity between peaks 6 and 7 from
1.31 to 1.99. The increase in selectivity is due to changes in the partition coefficients as a
result of the increased polarity of the mobile phase. As the mobile phase becomes more
polar, the analytes partition more into the stationary phase, and they are significantly
retained. As a consequence of the latter, the migration times are longer, the resolution is
higher, and the efficiency is lower. A slight increase in electroosmotic mobility, at higher
concentrations of ACN, was also observed from 1.57x10-4 to 3.56x10-4 cm2V-1s-1, probably
due to an increase in the ratio of the dielectric constant to buffer viscosity.
2.3.3 Effect of Applied Voltage
The effect of the applied voltage on the CEC separation of benzodiazepines was
then investigated using a mobile phase of 10 mM Tris (pH 8)-ACN (60:40). As expected,
retention times decreased when a higher voltage was applied. Figure 2.4 demonstrates the
electrochromatograms obtained when 30 kV, 20 kV and 15 kV were applied. At 30 kV, the
analytes elute faster with lower resolution and higher efficiency. Although the resolution
74
between analytes 2 and 3, and 5 and 6 is higher at 15 kV, the total separation time is
longer. Based on these results, 20 kV was applied to further optimize the separation
conditions.
1,2 3,4
5 6 7
3,4 2 1 6 5 7
1,2 5 6 3 4 7
min20 40 60 80 100 120
mAU
0 20 40 60 80
min20 40 60 80 100 120
mAU
0 5
10 15 20 25
min20 40 60 80 100 120
mAU
-20 0
20 40
6 0 % A C N
7 0 % M e O H
4 5 % T H F
Figure 2.2 Effect of the nature of organic modifier on the CEC separation of benzodiazepines. Conditions: C18 stationary phase; 40 cm packed x 100 µm i.d.; electrolyte, 10 mM Tris (pH 8)-ACN (40:60), 10 mM Tris (pH 8)-MeOH (30:70), 10 mM Tris (pH 8)-THF (55:45); applied voltage, 30 kV; electrokinetic injection, 15 kV for 5 s; temperature, 25 °C; UV detection, 220 nm.
75
1,2,3,4 5 6 7 1,2 3,4 7 5 6 1 2 5,6 3 4 7
1 2 3 4
5 6 7
min20 40 60 80
mAU
0 20 40 60
min20 40 60 80
mAU
0 20 40 60 80
min20 40 60 80
mAU
0 10 20 30 40
min20 40 60 80
mAU
0 5
10
A C N = 7 0 %
A C N = 6 0 %
A C N = 5 0 %
A C N = 4 0 %
Figure 2.3 Effect of mobile-phase composition on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the mobile phase composition (ACN-Tris) was varied.
2.3.4 Effect of Tris Concentration
The influence of the ionic strength of the mobile phase was also evaluated using 10
mM, 20 mM, and 30 mM Tris (pH 8)-ACN (60:40). At a constant ACN content, as the
ionic strength increased from 10 mM to 30 mM, the retention time of the analytes
increased (Figure 2.5). In addition, the electroosmotic mobility decreased from 1.76x10-4
to 1.30x10-4 cm2V-1s-1 with increasing Tris concentration due to the interactions of Tris
76
with the silica interface, which reduces the zeta potential. As mentioned in Chapter 1, the
electroosmotic mobility in packed-CEC is given by:
η
εεσµ
EcF
RTro
EOF
2/1
32
= (2.2)
where σ , which is proportional to zeta potential, is the charge density at the surface, oε is
the permittivity of vacuum (8.85 x 10-12 C2N-1µ-2), rε is the dielectric constant of the
mobile phase, R is the gas constant, T is temperature, c is the concentration of the
electrolyte, F is Faraday’s constant, η is the viscosity of the mobile phase, and E is the
electric field strength. According to the above equation, the electroosmotic mobility is
inversely proportional to the square root of the buffer concentration. Resolution also
increases upon increasing ionic strength due to improved stacking during electrokinetic
injection. An increase in ionic strength from 10 mM to 30 mM increased the resolution
between the analytes 5 and 6 from 1.04 to 1.39. Therefore, 30 mM Tris was used to further
optimize the separation.
2.3.5 Effect of Column Temperature
For this component of our study the temperature was varied from 45 °C to 15 °C,
and the binary mixture 30 mM Tris (pH 8)-ACN (60:40) was used as the mobile phase. As
in CE, the electroosmotic mobility increases upon increasing temperature in CEC, due to
the decrease in viscosity of aqueous-organic solvent system. When the temperature
increased from 15 °C to 45 °C, the electroosmotic mobility increased from 1.24x10-4 to
1.82x10-4 cm2V-1s-1. As is also typical for liquid chromatography, retention factors,
retention times, and resolution decrease at higher temperature. The effects of these
77
parameters by temperature variation are shown in Figure 2.6. It is also shown that by
decreasing the temperature to 15 °C all benzodiazepines were baseline resolved.
1 23 4
6 7 5 1 2 3 4 5 6 7 1 2 3 4 5 6 7
min20 40 60 80 100 120
mAU
-2.5 0 2.5 5 7.5 10 12.5
min20 40 60 80 100 120
mAU
-10 0
10 20 30
min20 40 60 80 100 120
mAU
-5 0 5
10 15 20
V = 3 0 k V
V = 2 0 k V
V = 1 5 k V
Figure 2.4 Effect of applied voltage on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the applied voltage was varied; electrolyte, 10 mM Tris (pH 8)-ACN (60:40).
78
12 3 4 5 6 7 1 2 3 4 5 6 7 1 2 3 4 5 6 7
min20 40 60 80 100 120
mAU
-10 0
10 20 30
min20 40 60 80 100 120
mAU
-10 0
10 20 30
min20 40 60 80 100 120
mAU
-10 0
10 20 30 40
1 0 m M T r i s
2 0 m M T r i s
3 0 m M T r i s
Figure 2.5 Effect of Tris concentration on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the Tris buffer concentration was varied; electrolyte, Tris (pH 8)-ACN (60:40); applied voltage, 20 kV.
2.3.6 CEC Separation of Drugs from a Urine Sample
The sample used in this study was a urine calibrator that contained 1000 ng/ml
oxazepam and other drugs, such as acetaminophen, amphetamines, imipramine, morphine,
cocaine, etc. Without any sample preparation the urine sample was injected into the CEC
capillary and the optimum conditions were applied. Figure 2.7a illustrates that the trace
79
amount of oxazepam was able to be detected and separated from the other drugs. The urine
sample was injected at 15 kV for 5 s. We also proved the presence of oxazepam in the
urine sample by spiking with the standard analyte (Figure 2.7b.). For this experiment, the
urine sample was injected at 20 kV for 10 s, and the standard analyte at 15 kV for 5 s.
2,3 1 4 5,6 7
1 2 3 4 5,6 7
1 2 3 4 5 6 7 1 2 3 4 5 6 7
min20 40 60 80 100
mAU
0 20 40
min20 40 60 80 100
mAU
-20 -10 0 10 20
min20 40 60 80 100
mAU
-10 0 10 20 30
min20 40 60 80 100
mAU
-10 0
10 20
T = 4 5° C
T = 3 5° C
T = 2 5° C
T = 1 5° C
Figure 2.6 Effect of column temperature on the CEC separation of benzodiazepines. Separation conditions are the same as in Figure 2.2, except the temperature was varied; electrolyte, 30 mM Tris (pH 8)-ACN (60:40); applied voltage, 20 kV.
80
? ? ?
1
min10 20 30 40 50
mAU
-10 0
10 20 30 40 50 60
(a)
(b) ? 1 ? ?
min0 10 20 30 40 50
mAU
0
200
400
600
800
Figure 2.7 CEC separation of drugs from a urine sample. (a) Without spiking. Conditions: C18 stationary phase; 40 cm packed x 100 µm i.d.; electrolyte, 30 mM Tris (pH8)-ACN (60:40); applied voltage, 20 kV; electrokinetic injection, 15 kV for 5 s; temperature, 15 °C; UV detection, 220 nm. (b) With spiking. Separation conditions are the same as above, except electrokinetic injection (urine sample), 20 kV for 10 s; electrokinetic injection (2 mg/ml standard oxazepam), 15 kV for 5 s.
81
2.4 Conclusion
In this study, we have developed a new CEC method for the baseline resolution of
benzodiazepines. The optimized method proved to be effective in separating and
identifying oxazepam in urine samples that contain various concentrations of other drugs.
We conclude that this CEC method is promising for determining benzodiazepines in
forensic and clinical drug analysis without any sample preparation. We have also been able
to understand how several parameters affect the electrophoretic and separation mechanisms
of hydrophobic analytes on an ODS stationary phase. The volume fraction of ACN and the
temperature are inversely proportional to migration time and resolution. However,
electroosmotic mobility increases upon increasing the volume fraction of ACN or the
temperature. In contrast, the ionic strength of the electrolyte is proportional to the
migration time and resolution, and inversely proportional to electroosmotic mobility.
Finally, higher resolutions were obtained at 15°C when a binary mixture of 30 mM Tris
(pH 8)-ACN (60:40) was used.
2.5 References
1. Linnoila, M. In Molecular Biology to Clinical Practice; Costa, E., Ed.; Raven Press: New York, NY, 1983, 267-278.
2. Gill, R.; Law, B.; Gibbs, J. P. J. Chromatogr. 1986, 356, 37.
3. Sternbach, L. H.; Randall, L. O.; Banziger, R.; Lehr, H. In Drugs affecting central
nervous system; Nutley, N. J., Ed.; Marcel Dekker, Inc.: New York, NY, 1968, 237-264.
4. Sioufi, A.; Dubois, J. P. J. Chromatogr. 1990, 531, 459.
5. Doble, A; Martin, I. L. Trends in Pharmacological Sciences 1992, 13, 76.
6. Drummer, O. H. J. Chromatogr. B 1998, 713, 201.
82
7. Smyth, W. F.; McClean, S. Electrophoresis 1998, 19, 2870.
8. Brettell, T. A.; Inman, K.; Rudin, N.; Saferstein, R. Anal. Chem. 2001, 73, 2735.
9. Tomita, M.; Okuyama, T. J. Chromatogr. B 1996, 678, 331.
10. Douse, J. M. F. J. Chromatogr. 1984, 301, 137.
11. Drouet-Coassolo, C.; Aubert, C.; Coassolo, P.; Cano, J. P. J. Chromatogr. 1989, 487, 295.
12. Evenson, M. A.; Wiktorowicz, J. E. Clin. Chem. 1992, 38, 1847.
13. Imazawa, M.; Hatanaka, Y. Journal of Pharmaceutical and Biomedical Analysis
1997, 15, 1503.
14. De Silva, J. A.; Bekersky, I.; Puglisi, C. V.; Brooks, M. A.; Weinfeld, R. E. Anal. Chem. 1976, 48, 10.
15. Dixon, W. R.; Earley, J.; Postma, E. J. Pharm. Sci. 1975, 64, 937.
16. Lloyd, J. B. F.; Parry, D. A. J. Chromatogr. 1988, 449, 281.
17. Mascher, H.; Nitsche, V.; Schutz, H. J. Chromatogr. 1984, 306, 231.
18. Klockowski, P. M.; Levy, G. J. Chromatogr. 1987, 422, 334.
19. Vree, T. B.; Baars, A. M.; Hekster, Y. A.; Van der Kleijn, E. J. Chromatogr. 1981,
224, 519.
20. Minder, E. I.; Schaubhut, R.; Minder, C. E.; Vonderschmitt, D. J. J. Chromatogr. 1987, 419, 135.
21. Aakerman, K. K.; Jolkkonen, J.; Parviainen, M.; Penttilae, I. Clin. Chem. 1996, 42,
1412.
22. Schafroth, M.; Thormann, W.; Allemann, D. Electrophoresis 1994, 15, 72.
23. Tomita, M.; Okuyama, T.; Sato, S.; Ishizu, H. J. Chromatogr. 1993, 621, 249.
24. Wernly, P.; Thormann, W. Anal. Chem. 1992, 64, 2155.
25. Inoue, T.; Niwaguchi, T. J. Chromatogr. 1985, 339, 163.
26. Mule, S. J.; Casella, G. A. J. Anal. Toxicol. 1989, 13, 179.
83
27. Maurer, H.; Pfleger, K. J. Chromatogr. 1987, 422, 85.
28. Yegles, M.; Mersch, F.; Wennig, R. Forensic Science International 1997, 84, 211.
29. Tagliaro, F.; Smyth, W. F.; Turrina, S.; Deyl, Z.; Marigo, M. Forensic Science International 1995, 70, 93.
30. Brooks, M. A.; D’Arconte, L.; Hackman, M. R.; De Silva, J. A. F. J. Anal. Toxicol.
1997, 1, 179.
31. Brooks, M. A.; Bruno, J. J. B.; De Silva, J. A. F.; Hackman, M. R. Anal. Chim. Acta 1975, 74, 367.
32. Werner, I. A.; Altorfer, H.; Perlia, X. Chromatographia 1990, 30, 255.
33. Smith, R. M.; Sanagi, M. M. J. Chromatogr. 1989, 483, 51.
34. Valko, K.; Olajos, S.; Cserhati, T. J. Chromatogr. 1990, 499, 361.
35. Mura, P.; Piriou, A.; Fraillon, P.; Papet, Y.; Reiss, D. J. Chromatogr. 1987, 416,
303.
36. Guillaume, Y.; Guinchard, C. J. Liq. Chromatogr. 1993, 16, 3457.
37. Guillaume, Y.; Guinchard, C. J. Liq. Chromatogr. 1994, 17, 1443.
38. Choma, I.; Dawidowicz, A. L.; Lodkowski, R. J. Chromatogr. 1992, 600, 109.
39. Renou-Gonnord, M. F.; David, K. J. Chromatogr. A 1996, 735, 249.
40. Nishi, H.; Tsumagari, N.; Kakimoto, T.; Terabe, S. J. Chromatogr. 1989, 465, 331.
41. Terabe, S.; Otsuka, K.; Ichikawa, K.; Tsuchiya, A.; Ando, T. Anal. Chem. 1984, 56, 111.
42. Boonkerd, S.; Detaevernier, M. R.; Vindevogel, J.; Michotte, Y. J. Chromatogr. A
1996, 756, 279.
43. Yan, C.; Dadoo, R.; Zhao, H.; Zare, R. N.; Rakesraw, D. J. Anal. Chem. 1995, 67, 2026.
44. Catabay, A. P.; Sawada, H.; Jinno, K.; Pesek, J. J.; Matyska, M. T. J. Capillary
Electrophor. 1998, 5, 89.
45. Cahours, X.; Morin, P.; Dreux, M. J. Chromatogr. A 1999, 845, 203.
84
46. Thiam, S.; Shamsi, S. A.; Henry, C. W.; Robinson, J. W.; Warner, I. M. Anal. Chem. 2000, 72, 2541.
47. Henry, C. W.; McCarroll, M. E.; Warner, I. M. J. Chromatogr. A 2001, 905, 319.
48. Moffatt, F.; Cooper, P. A.; Jessop, K. M. J. Chromatogr. A 1999, 855, 215.
49. Smith, N. W.; Evans, M. B. Chromatographia 1994, 38, 649.
50. Reilly, J.; Saeed, M. J. Chromatogr. A 1998, 829, 175.
51. Wang, J.; Schaufelberger, D. E.; Guzman, N. A. J. Chromatogr. Sci. 1998, 36, 155.
52. Jinno, K.; Sawada, H.; Catabay, A. P.; Watanabe, H.; Haji Sabli, N. B.; Pesek, J. J.;
Matyska, M. T. J. Chromatogr. A 2000, 887, 479.
53. Matyska, M. T.; Pesek, J. J.; Katrekar, A. Anal. Chem. 1999, 71, 5508.
54. Knox, J. H.; Grant, I. H. Chromatographia 1987, 24, 135.
55. Knox, J. H.; Grant, I. H. Chromatographia 1991, 32, 317.
56. Schoenmakers, P. J.; Billiet, H. A. H.; De Galan, L. J. Chromatogr. 1979, 185, 179.
57. Schoenmakers, P. J.; Billiet, H. A. H.; De Galan, L. J. Chromatogr. 1981, 218, 261.
58. Dittmann, M. M.; Rozing, G. P. J. Chromatogr. A 1996, 744, 63.
85
CHAPTER 3.
ANALYTICAL SEPARATIONS USING POLYMERIC SURFACTANTS IN OPEN-TUBULAR CAPILLARY ELECTROCHROMATOGRAPHY
3.1 Introduction
Capillary electrochromatography (CEC) is a hybrid electroseparation technique that
couples the selectivity of high performance liquid chromatography (HPLC) and the
separation efficiency of capillary electrophoresis (CE) [1-5]. CEC also provides high
resolution, short analysis time, smaller sample and buffer consumption, and efficiencies
five to ten times higher than HPLC. The separation in CEC is based upon the
electrophoretic mobility of the solutes and their partitioning between the stationary and
mobile phases.
In the development of CEC, both packed and open-tubular column configurations
have been reported [1-10]. The packed mode of CEC utilizes a fused-silica capillary with a
typical internal diameter of 50-100 µm. This capillary is packed with a typical HPLC
stationary phase such as an octadecyl silica (ODS) stationary phase [2, 4]. However, there
are several problems that need to be solved in order for packed-CEC to be a viable
alternative to either CE or HPLC. The limitations of conventional CEC include the
necessity to fabricate frits, the tendency that packed capillaries have to form bubbles
around the packing material or at the frit, the difficult packing procedure, and the difficult
separation of basic compounds. The problems mentioned above are discussed in detail in
Chapter 1.
Open-tubular CEC (OT-CEC) is an alternative approach to packed-CEC [9]. None
of the problems mentioned above are likely to be encountered in an open-tubular format. In
86
this CEC format, a stable coating needs to be constructed on the inner walls of the capillary
in order to provide efficient chromatographic separations and reproducible EOF [10]. The
most commonly used approaches to wall coatings for modifying the capillary include: (i)
dynamic coating performed by adding the cationic or neutral modifier to the electrolytes
[11, 12]; (ii) adsorbed cationic modifier on the capillary wall by physical adsorption [13-
16]; and (iii) fixation of the hydrophilic layer by covalent bonding and/or cross linking
[17-22]. Harrell et al. [23] achieved a baseline separation of seven tricyclic antidepressants
by use of a novel nonionic micelle polymer, poly (n-undecyl-α-D-glucopyranoside) as a
dynamic coating. However, dynamic coating is known to cause problems when CE is
coupled to mass spectrometry (CE/MS). In addition, the presence of the nonvolatile buffer
constituents may deteriorate the ionization of the analytes [24, 25]. Although physical
adsorption has a simple and rapid coating procedure and good reproducibility, it has been
shown to have a short lifetime and limited pH range [24, 26]. In contrast, some of the
covalent bonding and/or cross-linking have a long lifetime, but require a more complicated
coating procedure [24, 26]. Obviously, an ideal coating procedure would be one that is
both simple and stable.
In this chapter, an alternative to covalent linking of a polymer to silica beads is
explored. In our approach, the polymeric surfactant poly (sodium N-undecanoyl-L-
glycinate), poly (L-SUG), is used in a simple coating procedure, which involves a layer-by-
layer deposition process [27, 28]. This coating is a polyelectrolyte multilayer (PEM), and it
is constructed in situ by alternating rinses of positively and negatively charged polymers
[29-33]. Via electrostatic forces, a layer of polymer adds to the oppositely charged surface,
reversing the surface charge and priming the film for the addition of the next layer. Such
87
coatings have been found to be robust, and thus, highly resistant to charge and
deterioration during use [24, 26, 32]. The advantages of our PEM coating are two-fold.
First, since the polymeric surfactant is coated electrostatically onto the capillary, less
consumption of the reagent is required. Second, with the polymeric surfactant coated on
the capillary, there is less detection interference with the analyte of interest, which in turn
makes the system more amenable to coupling with mass spectrometry or other detectors
where the polymeric surfactant reagent interferes.
This PEM coating approach used for fabricating columns for use in OT-CEC is
described below. The performance of the modified capillaries as a separation medium is
evaluated by use of seven benzodiazepines as analytes. The coating was found to be
remarkably stable with excellent performance for more than 200 runs.
3.2 Experimental
3.2.1 Apparatus and Conditions
Separations were performed on a Beckman P/ACE MDQ capillary electrophoresis
system with UV detection (Fullerton, CA). The fused-silica capillary, 57 cm (50 cm
effective length) x 50 µm i.d., was purchased from Polymicro Technologies (Phoenix, AZ)
and mounted in a Beckman capillary cartridge. Unless stated otherwise, the cartridge
temperature was maintained at 25 °C by use of liquid coolant. UV detection was performed
at 214 nm and the samples were injected by pressure (0.1 psi; 1 psi=6894.76 Pa) for 1 sec.
3.2.2 Reagents and Chemicals
Flunitrazepam, temazepam, diazepam, oxazepam, lorazepam, clonazepam and
nitrazepam were purchased from Sigma Chemical Company (St Louis, MO). The
structures of the analytes used in this study are shown in Figure 2.1 (Chapter 2). However,
88
the elution order of benzodiazepines changes to flunitrazepam1 <temazepam2 <diazepam3
<oxazepam4 <lorazepam5 <clonazepam6 <nitrazepam7. Sodium phosphate (Na2HPO4 and
NaH2PO4), hydrochloric acid (HCl) and sodium chloride (NaCl) were all obtained from
Fisher Scientific (Fair Lawn, NJ). Poly (diallyldimethylammonium chloride), PDADMAC
(Mw=200,000-350,000) was obtained from Aldrich (Milwankee, WI). Other chemicals,
including L-glycine, undecylenic acid and N-hydroxysuccinimide, were also purchased
from Sigma.
3.2.3 Sample and Buffer Preparation
Analytical standard benzodiazepine stock solutions were prepared in methanol-
water (1:1) at concentrations of about 0.15 mg/ml each. A buffer solution of 50 mM
Na2HPO4 was prepared by dissolving the appropriate amount of Na2HPO4 in 10 ml of
deionized water. The solution was filtered using a polypropylene nylon filter with 0.45 µm
pore size and sonicated for 15 min before use.
3.2.4 Synthesis of Monomeric and Polymeric Surfactant
The surfactant monomer of sodium N-undecenoyl-L-glycinate, mono (L-SUG), was
synthesized from the N-hydroxysuccinimide ester of undecylenic acid according to a
previously reported procedure [34]. A 100 mM sodium salt solution of the monomer was
then polymerized by use of 60Co-γ radiation. After irradiation, the polymer was dialyzed by
use of a 2000 molecular mass cut-off and then lyophilized to obtain the dry product.
Structures of the monomeric and polymeric surfactants are illustrated in Figure 3.1.
3.2.5 Procedure for Polyelectrolyte Multilayer (PEM) Coating
PEM coating was achieved by deposition of the polymer solutions using the rinse
function on the Beckman CE system. Each polymer deposition solution contained 0.5%
89
(w/v) polymer in 0.2 M aqueous NaCl solution. It was observed that the addition of NaCl
to the polymer solution resulted in enhanced thickness for each polyelectrolyte layer [32].
The capillary was conditioned before coating using a 5-min rinse of water in order to
remove any contaminants originating from the capillary drawing process. The column was
then conditioned with 1 M NaOH for 60 min. Pure deionized water was flushed through
the capillary for 15 more min. The first monolayer of polymer (PDADMAC) was
deposited by rinsing the solution of the cationic polymer through the capillary for 20 min
followed by a 5-min water rinse. All other polymer depositions were done with 5-min
rinses followed by 5-min water rinses. A diagrammatic scheme of the PEM-coated
capillary is shown in Figure 3.2. This diagram is not provided to give an actual structural
representation of the bilayer, but only to represent the order of polymer deposition. The
multilayer coatings used for the separation of benzodiazepines and the reproducibility
studies consisted of ten layer pairs (a layer pair is a layer of cationic polymer plus a layer
of anionic polymer; also termed a bilayer). The capillary was then flushed with buffer until
a stable current was achieved. The columns were conditioned with buffer for 2 min
between injections.
3.3 Results and Discussion
3.3.1 Endurance of PEM Coating
An important aspect of this approach, which must be considered, is the lifetime of
the stationary phase. Thus, we examined the stability of our coating by use of the following
procedure. A 10-layer pair multilayer was constructed, and nearly 50 separations were
performed within 5 days. Each separation was done with applied voltages of 15 kV to 30
kV at 25 °C. The electrolyte concentration was 50 mM of Na2HPO4 and a pH range of 9.2-
90
11.0 was used. After these experiments, the capillary was removed from the instrument,
and the tips of the column were placed in water vials for one week. When the capillary was
placed back in the instrument, 50 replicate runs were performed using an applied voltage
of 20 kV, a temperature of 25 °C, and a 50-mM phosphate buffer (pH 9.2). The capillary
was again removed from the instrument and the tips were placed in water vials for an
additional week, after which the column was placed back into the instrument and more
than 100 runs were performed. In these studies, both the applied voltage and the
temperature were varied from 15 kV to 30 kV and from 15 °C to 35 °C, respectively.
Therefore, the aggregate performance of the PEM-coated capillary was evaluated for more
than 200 runs.
CO
NH
C O
Na +O-
CONH
C ONa+O-
CO NHCO
Na+O-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHN
CO-ONa+
CO
HNC
O
Na+O-
CO
HNC
O
Na+O-
CO
NHC
ONa+O-
(a) (b)
Figure 3.1 Structures of the (a) monomeric SUG and (b) polymeric SUG.
91
SiO- SiO- SiO- SiO- SiO- SiO- SiO- SiO- SiO-
N+ N+ N+ N+ N+ N+ N+ N+ N+ N+N+N+ N+
N+N+N+N+N+N+N+N+N+N+ N+N+N+
Capillary wall, silanol groups
Cationic polymer, PDADMAC
Anionic polymer, poly(L-SUG) CO
NH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
COHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHC
O
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-
CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
COHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHC
O
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-
CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHC
O
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O
-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-
CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHC
O
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO- CONH
C O+NaO-
CO NHC O
+NaO-
CO
NHCO
-ONa+
CO
NH
CO
-ONa+
CO
HNC
O-ONa+
C OHN
CO -ONa+
C OHNCO-ONa+
CO
HNC
O
+NaO-
CO
HNC
O+NaO-
CO
NHC
O+NaO-
Figure 3.2 Scheme of the PEM-coated capillary.
3.3.2 Stability of PEM Coating
Another important factor to consider when using PEM-coated capillaries is the
stability of the capillary surface, especially after exposure to solutions with extreme pH
values. To evaluate this parameter, two more phosphate buffers of pH 11.0 and 3.0 were
prepared. First, 30 replicate runs were performed with 50 mM phosphate buffer (pH 9.2).
The 10th run (Figure 3.3a) yielded an electroosmotic mobility of 2.39x10-3 cm2V-1s-1. The
capillary was then flushed with 50 mM phosphate buffer (pH 11.0) for 100 min and 50
mM phosphate buffer (pH 9.2) for 30 min. One of the electropherograms obtained after the
exposure to pH 11.0 (Figure 3.3b) gave an electroosmotic mobility of 2.37x10-3 cm2V-1s-1.
After this, the same procedure was followed with the 50 mM phosphate buffer (pH 3.0).
One of the runs that was performed after the last exposure (Figure 3.3c) yielded an
electroosmotic mobility of 2.37x10-3 cm2V-1s-1. Therefore, the PEM coating was
demonstrated to have extraordinary stability under extreme values of pH.
92
3.3.3 Reproducibilities
Reproducibility of the PEM coating is also an important consideration. The
reproducibilities were evaluated by computing the relative standard deviations (RSDs) [36]
of the EOF, which are reported in Table 3.1. The run-to-run RSD was obtained from 50
consecutive electrophoresis runs; both the day-to-day and capillary-to-capillary RSDs were
obtained by use of five replicate analyses; the week-to-week RSD was obtained by use of
three replicate analyses. All RSDs of the EOF were below 1%, and thus, very good
reproducibilities were observed.
3.3.4 Voltage Study
The PEM-coated capillary was applied to the separation of seven benzodiazepines.
The influence of the applied voltage on the efficiency, resolution, and analysis time of the
benzodiazepines was evaluated using a mobile phase of 50 mM Na2HPO4 at 25 °C. As
expected, a higher voltage decreased the retention times. At 30 kV, the analytes eluted
faster with higher efficiency and lower resolution. In contrast, at 15 kV, the migration
times were longer, the resolution was higher, and the efficiency was lower. However, an
applied voltage of 15 kV did not have a major impact on analyte resolution, compared to
the electropherogram obtained when a 20 kV voltage was applied (Figure 3.4).
3.3.5 Temperature Study
The effect of temperature on the separation of benzodiazepines was also studied.
The temperature for this study was varied from 35 °C to 15 °C. As shown in Figure 3.5, the
retention time decreased at higher temperature, and peak efficiency decreased at lower
temperature. In addition, electroosmotic mobility increased when temperature increased,
likely due to a decrease in electrolyte viscosity.
93
a
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
(a) run 10
(b) after exposure to pH 11.0 (run 40)
(c) after exposure to pH 3.0 (run 110)
123 4 5 6 7
Figure 3.3 Stability studies of PEM coating. Conditions: 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SUG) with 0.2 M NaCl; pressure injection, 0.1 psi for 1 s; electrolyte, 50 mM Na2HPO4 (pH 9.2); applied voltage, 20 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm.
94
Table 3.1 Reproducibilities of PEM capillary coating. Conditions: same as Figure 3.3.
RSD (%)EOF Average (min)
0.789.990run-to-run
day-to-day
capillary-to-capillary
week-to-week
9.982
9.930
9.950
0.81
0.86
0.93
RSD (%)EOF Average (min)
0.789.990run-to-run
day-to-day
capillary-to-capillary
week-to-week
9.982
9.930
9.950
0.81
0.86
0.93
3.3.6 Comparison Between Monomeric and Polymeric Surfactants
Another important consideration for this study is whether molecular micelles are
needed to form an effective and stable PEM, or can the same be achieved by use of
monomeric surfactants. In an effort to compare the chromatographic performance of mono
(L-SUG) and poly (L-SUG) for the separation of hydrophobic analytes, benzodiazepines
were used as test solutes. The separation of benzodiazepines using 50 mM Na2HPO4 (pH
9.2) as the electrolyte and 0.5% (w/v) poly (L-SUG) as the anionic polyelectrolyte for the
construction of the PEM coating is shown in Figure 3.6a. Figure 3.6b is the
electropherogram of the benzodiazepines under the same conditions as in Figure 3.6a.
However, the anionic surfactant used for PEM coating construction in this figure is the
monomeric (nonpolymerized) surfactant at the concentration of 0.5% (w/v) (19 mM).
Almost no separation is noted, even though the monomeric surfactant concentration is
significantly above the normal CMC of the nonpolymerized surfactant (7 mM). Thus, it is
95
clear from this study that the molecular micelle allows better discrimination of the
hydrophobic analytes than the conventional micelle.
In a normal (nonpolymerized) micellar system, the dynamic equilibrium that exists
between the monomers and micellar aggregates, has been demonstrated to be a
disadvantage for separations.33 This dynamic equilibrium will likely reduce the stability of
the PEM coating in OT-CEC. In such a case, poor analyte separation will be observed as
seen here. In contrast, polymeric micelles do not have such problems because the covalent
bonds formed between monomers eliminate dynamic equilibrium. Thus, the coating that is
produced will be more stable as is observed in this study.
3.4 Conclusion
A stable modified capillary has been developed by use of a simple PEM coating
procedure employing a molecular micelle. Excellent run-to-run, day-to-day, week-to-week
and capillary-to-capillary reproducibilities in separation were observed since the RSD
values of electroosmotic flow were below 1% in all cases. In addition, the PEM-coated
capillaries exhibited high stability against extreme pH values. The stability of the
capillaries allowed us to perform over 200 runs. This approach also shows that highly
efficient and reproducible peaks can be obtained from stationary phases prepared by such a
simple procedure. In addition, the chromatographic performance of the monomeric form of
the molecular micelle was compared for the separation of benzodiazepines. This study
confirmed that the polymeric surfactant allows better discrimination of hydrophobic
analytes than the nonpolymerized surfactant. We conclude that this method is a promising
alternative to conventional CEC and should prove very useful in both chiral and achiral
separations.
96
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
30 kV
15 kV
20 kV
123 4 5 6 7
Figure 3.4 Effect of applied voltage on the OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except applied voltage was varied.
97
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34
AU
0.000
0.001
0.002
0.003
0.004
35°°°°C
25°°°°C
15°°°°C
123 4 5 6 7
Figure 3.5 Effect of temperature on the OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except the temperature was varied.
98
Minutes0 2 4 6 8 10 12 14 16 18 20
AU
0.000
0.002
0.004
0.006
0.008
0.010
Minutes0 2 4 6 8 10 12 14 16 18 20
AU
0.000
0.002
0.004
0.006
0.008
0.010
(a) poly(L-SUG)
(b) mono(L-SUG)
123 4 5 6 7
Figure 3.6 Comparison between monomeric and polymeric surfactants for OT-CEC separation of benzodiazepines. Conditions: same as Figure 3.3, except applied voltage, 30 kV. (a) 0.5% (w/v) poly (L-SUG) (b) 0.5% (w/v) mono (L-SUG).
3.5 References
1. Liu, Z.; Zou, H.; Ye, M.; Ni, J.; Zhang, Y. Electrophoresis 1999, 20, 2891.
2. Henry, C. W.; McCarroll, M. E.; Warner, I. M. J. Chromatogr. A 2001, 905, 319.
3. Ye, M.; Zou, H.; Liu, Z.; Ni, J.; Zhang, Y. Anal. Chem. 2000, 72, 616.
4. Thiam, S.; Shamsi, S. A.; Henry, C. W.; Robinson, J. W.; Warner, I. M. Anal. Chem. 2000, 72, 2541.
5. Jorgenson, J. W.; Lukacs, K. D. Anal. Chem. 1981, 53, 1298.
99
6. Jinno, K.; Sawada, H.; Catabay, A. P.; Hiroshi, W.; Sabli, N. B. H.; Pesek, J. J.; Matyska, M. T. J. Chromatogr. A 2000, 887, 479.
7. We, J.; Huang, P.; Li, M. X.; Qian, M. G.; Lubman, D. M. Anal. Chem. 1997, 69,
320.
8. Dulay, M. T.; Quirino, J. P.; Bennett, B. D.; Kato, M.; Zare, R. N. Anal. Chem. 2001, 73, 3921.
9. Matyska, M. T.; Pesek, J. J.; Katrekar, A. Anal. Chem. 1999, 71, 5508.
10. Hayes, J. D.; Malik, A. Anal. Chem. 2001, 73, 987.
11. Gilges, M.; Kleemiss, M. H.; Schomburg, G. Anal. Chem. 1994, 66, 2038.
12. Cifuentes, A.; Poppe, H.; Kraak, J. C.; Erim, F. B. J. Chromatogr. B 1996, 681, 21.
13. Erim, F. B.; Cifuentes, A.; Poppe, H.; Kraak, J. C. J. Chromatogr. A 1995, 708,
356.
14. Chiu, R. W.; Jimenez, J. C.; Monnig, C. A. Anal. Chim. Acta 1995, 307, 193.
15. Preisler, J.; Yeung, E. S. Anal. Chem. 1996, 68, 2885.
16. Cordova, E.; Gao, J.; Whitesides, G. M. Anal. Chem. 1997, 69, 1370.
17. Towns, J. K.; Regnier, F. E. J. Chromatogr. 1990, 516, 69.
18. Figeys, D.; Aebersold, R. J. Chromatogr. B 1997, 695, 163.
19. Bruin, G. J. M.; Chang, J. P.; Kuhlman, R. H.; Zegers, K.; Kraak, J. C.; Poppe, H. J. Chromatogr. 1989, 471, 429.
20. Hjerten, S.; Johansson, M. K. J. Chromatogr. 1991, 550, 811.
21. Cobb, K. A.; Dolnik, V.; Novotony, M. Anal. Chem. 1990, 62, 2478.
22. Huang, X.; Horvath, C. J. Chromatogr. A 1997, 788, 155.
23. Harrell, C. W.; Dey, J.; Shamsi, S. A.; Foley, J. P.; Warner, I. M. Electrophoresis
1998, 19, 712.
24. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 5272.
25. Niessen, W. M. A.; Tjaden, U. R.; Geef, J. J. Chromatogr. 1993, 636, 3.
100
26. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 2254.
27. Decher, G.; Schmitt, J. Prog. Colloid Polym. Sci. 1992, 89, 160.
28. Decher, G. Science 1997, 277, 1232.
29. Schlenoff, J. B.; Ly, H.; Li, M. J. Am. Chem. Soc. 1998, 120, 7626.
30. Laurent, D.; Schlenoff, J. B. Langmuir 1997, 13, 1552.
31. Schlenoff, J. B.; Li, M. Ber. Bunsen-Ges. Phys. Chem. 1996, 100, 943.
32. Graul, T. W.; Schlenoff, J. B. Anal. Chem. 1999, 71, 4007.
33. Schlenoff, J. B.; Dubas, S. T.; Farhat, T. Langmuir 2000, 16, 9968.
34. Wang, J.; Warner, I. M. Anal. Chem. 1994, 66, 3773.
35. Dittmann, M. M.; Rozing, G. P. J. Chromatogr. A 1996, 744, 63.
36. Skoog, D. A.; Holler, J. F.; Nieman, T. A. In Principles of Instrumental Analysis, 5th ed.; Sherman, M., Bortel, J., Messina, F., Eds.; Harcourt Brace College Publishers: Orlando, FL, 1998.
101
CHAPTER 4.
CHIRAL SEPARATIONS USING POLYMERIC SURFACTANTS AND POLYELECTROLYTE MULTILAYERS IN OPEN-TUBULAR CAPILLARY
ELECTROCHROMATOGRAPHY
4.1 Introduction
The separation of chiral compounds has been of great importance in many
industries, particularly the pharmaceutical industry. This interest is due to the different
pharmakokinetic characteristics and pharmacological activities of each enantiomer in a
racemic drug [1, 2]. Thus, there has been a great demand for the development of analytical
techniques for the separation of chiral bioorganic molecules.
In recent years, different modes of capillary electrophoresis (CE) have proven to be
very efficient methods for the separation and analysis of enantiomeric compounds [3-5].
Micellar electrokinetic chromatography (MEKC), which is one of the most recent methods
employed for chiral separations, uses a pseudostationary phase that involves the addition of
a chiral micellar selector in the background electrolyte (BGE).
In 1994, Wang and Warner reported the use of a synthetic chiral polymeric
surfactant added to the BGE for chiral separations. These polymeric amino acid-based
surfactants offer several advantages over conventional micelles [6-9]. As stated in Chapter
1, the polymerization of the surfactant eliminates the dynamic equilibrium between
monomers and micelles. This, in turn, results in better chiral recognition and enantiomeric
resolution. In addition, the polymeric surfactant can be used at very low concentrations
since it does not depend on the critical micelle concentration. This usually results in higher
efficiencies and rapid analysis in CE.
102
Recently, the use of polymeric surfactants with two amino acid functional groups
(dipeptides) at the polar head region has attracted considerable attention [10]. Various
polymeric dipeptide chiral surfactants have been synthesized and used as chiral
discriminators in MEKC [11, 12]. So far, poly (sodium N-undecanoyl-L-leucyl-valinate),
poly (L-SULV), has shown the best chiral discrimination ability for a wide variety of drug
compounds [8]. One major drawback to using these polymeric surfactants as additives in
BGE is that they are ultimately flushed out, and a fresh BGE is required for each
separation. To circumvent this problem, alternatives to MEKC have been explored [13-19].
Open-tubular capillary electrochromatography (OT-CEC) is an alternative
approach to MEKC. It requires the construction of a stable coating on the inner walls of the
capillary. This, in turn, provides efficient chromatographic separations and reproducible
electroosmotic flow (EOF) [20].
In 1995, Erim et al. [13] investigated the performance of a polyethyleneimine (PEI)
layer as a coating for the separation of basic proteins and peptides. The PEI coating was
constructed by just filling the capillary with a solution containing high-molecular-mass PEI
and flushing the capillary after a certain time. Katayama et al. [14, 15] reported the use of a
new simple coating procedure that is called successive multiple ionic-polymer layer
coating. In their studies, they developed both anion-modified and cation-modified
capillaries for the separation of acidic and basic proteins, respectively. The anion-modified
capillary was prepared by first attaching the cationic polymer to the capillary wall, and
then the anionic polymer to the cationic polymer layer. The cation-modified capillary was
established by attaching the cationic polymer to the anionic polymer layer. In 1999, Graul
and Schlenoff [16] used poly (styrene sulfonate) as the anionic polymer and poly
103
(diallyldimethylammonium chloride) as the cationic polymer in a polyelectrolyte
multilayer (PEM) coating procedure, which involved an electrostatic layer-by-layer
deposition process [21, 22]. This coating was constructed in situ by alternating rinses of
positively and negatively charged polymers [23-30], and it was applied to the separation of
a series of basic proteins. In addition, in 2003, Rmaile and Schlenoff [17] demonstrated
that the use of optically active PEMs for chiral membrane separations allows very high
enantiomer permeation rates with encouraging selectivity. In our laboratory, a PEM
coating has been successfully employed as a separation medium for the achiral separation
of benzodiazepines (Chapter 1) and phenols [18, 19].
In this chapter, the PEM coating is applied to chiral separations by the use of the
anionic polymer poly (L-SULV) as the chiral discriminator. An examination of the
influence of several parameters that are used to obtain more efficient and more
reproducible chromatographic separations is also discussed. The PEM-coated capillary was
found to be remarkably robust with a performance of more than 230 runs.
4.2 Experimental
4.2.1 Apparatus and Conditions
All experiments were conducted using a Beckman P/ACE MDQ capillary
electrophoresis system with UV detection (Fullerton, CA). Fused-silica capillaries, 57 cm
(50 cm effective length) x 50 µm i.d., were purchased from Polymicro Technologies
(Phoenix, AZ). The applied voltage was 30 kV. The samples were injected by pressure.
1,1’-binaphthyl-2,2’-dihydrogenphosphate (BNP), 1,1’-bi-2-naphthol (BOH), secobarbital
and pentobarbital were injected at 0.5 psi (1 psi = 6894.76 Pa) for 3 s, and temazepam at
0.9 psi for 7 s. Unless stated otherwise, the temperature of the cartridge was maintained at
104
25 °C using a liquid coolant. UV detection was performed at 214 nm for BNP, BOH,
secobarbital and pentobarbital, and 220 nm for temazepam.
4.2.2 Reagents and Chemicals
The analytes BNP, BOH, secobarbital, pentobarbital and temazepam, and
tris(hydroxymethyl)aminomethane (Tris) were purchased from Sigma Chemical Co. (St.
Louis, MO). Sodium borate (Na2B4O7) and sodium chloride (NaCl) were obtained from
Fisher Scientific (Fair Lawn, NJ). Boric acid (H3BO3) was purchased from Alfa Products
(Danvers, MA), and sodium phosphate (Na2HPO4) from Mallinckrodt & Baker, Inc. (Paris,
KY). The ionic liquids 1-ethyl-3-methyl-1H-imidazolium hexafluorophosphate (1E-3MI-
HFP) and 1-butyl-3-methylimidazolium tetrafluoroborate (1B-3MI-TFB) were purchased
from Aldrich Chemical Co. (Milwaukee, WI) and Chemada Fine Chemicals Ltd. (Nir
Itzhak, D. N. HaNegev 85455, Israel), respectively. The dipeptide L-leucylvalinate,
undecylenic acid, and N-hydroxysuccinimide were purchased from Sigma. Poly
(diallyldimethylammonium chloride) (PDADMAC; Mw = 200,000-350,000) was obtained
from Aldrich.
4.2.3 Sample and Background Electrolyte Preparation
The BGE at pH 10.0 consisted of 100 mM Tris and 10 mM Na2B4O7, the BGE at
pH 8.5 consisted of 25 mM Tris and 25 mM Na2B4O7, and the BGE at pH 7.2 consisted of
300 mM H3BO3 and 30 mM Na2HPO4. In all cases, the pH values were adjusted by using
either 1 M sodium hydroxide (NaOH) or 1 M hydrochloric acid (HCl). All solutions were
filtered using 0.45 µm polypropylene nylon filters and sonicated for 15 min before use.
The analytes BNP, BOH and secobarbital were dissolved in 50:50 methanol/water, and the
analytes pentobarbital and temazepam were dissolved in 80:20 methanol/water. The final
105
analyte concentrations were 0.1 mg/ml for BNP and BOH, 0.2 mg/ml for secobarbital, and
0.5 mg/ml for pentobarbital and temazepam.
4.2.4 Synthesis of Polymeric Surfactant
The monomeric surfactant of sodium N-undecenoyl-L-leucylvalinate was
synthesized from the N-hydroxysuccinimide ester of undecylenic acid using a procedure
reported by Wang and Warner [7]. A 100 mM sodium salt solution of the monomer was
then polymerized by use of 60Co-γ radiation. The structure of poly (L-SULV) is shown in
Figure 4.1.
HN
C
[CH-CH]x
CNH
CO-Na+
O
O
O
Figure 4.1 Structure of poly (L-SULV).
106
4.2.5 Procedure for Polyelectrolyte Multilayer Coating
PEM coatings were constructed in our laboratory according to a procedure
described in Chapter 3 [18]. The coatings were achieved by flushing the cationic and
anionic polymer deposition solutions through the capillaries. Polymer deposition solutions
contained 0.5% and 2.5% (w/v) polymer and 0.0-0.2 M NaCl or 0.01 M of the ionic liquid
1E-3MI-HFP or 1B-3MI-TFB. The capillary was first conditioned with water for 5 min,
with 1 M NaOH for 60 min, and then again with water for 15 min. The first layer of the
cationic polymer was deposited by flushing the PDADMAC solution through the capillary
for 5 min followed by a 5-min water rinse. All polymer depositions were done with 5 min
rinses followed by 5 min water rinses. The PEM coatings used for all the studies reported
here consisted of two layer pairs. However, for the chiral separations of the analytes
secobarbital and pentobarbital three layer pairs were constructed. Each coated capillary
was then flushed with the BGE until a stable current was achieved.
4.3 Results and Discussion
Several experimental parameters such as the type of additive in the polymer
deposition solutions, NaCl concentration, column temperature, and bilayer number were
studied to optimize the chiral separation of BNP. The influence of each parameter on chiral
separation was examined in detail as reported below. For the purpose of this work two
polymeric surfactants were examined: poly (sodium N-undecanoyl-L-valinate), poly (L-
SUV), and poly (L-SULV). The use of the first polymeric surfactant in the chiral
separation of BNP resulted in a single peak. Therefore, all studies were performed using
the polymeric surfactant poly (L-SULV).
107
4.3.1 Effect of Additives in Polymer Deposition Solutions
Each polymer deposition solution contained a polymer and an additive. NaCl was
one of the additives used for this study. It has been reported that the addition of NaCl to the
polymer solution resulted in enhanced thickness for each layer [16, 25]. Several studies
have also shown that salt concentration is approximately proportional to the thickness of
each polyelectrolyte layer [31-33].
The ionic liquids 1E-3MI-HFP and 1B-3MI-TFB were also used as additives in the
solutions used for deposition. Ionic liquids are liquid electrolytes composed entirely of ions
with melting points at ambient temperature. In recent years, there has been a growing
interest in ionic liquids due to their distinct properties. They are good solvents for several
organic, inorganic and polymeric materials, and they have excellent chemical and thermal
stability. They are also good electrical conductors, and nonvolatile [33-35]. With these
properties in mind, the ionic liquids were explored for possible enhanced separations.
Figures 4.2a, 4.2b and 4.2c demonstrate the chiral separation of BNP using 0.01 M
NaCl, and the ionic liquids 1E-3MI-HFP or 1B-3MI-TFB as additives in the polymer
solutions, respectively. The addition of either ionic liquid increased the resolution of BNP
from 0.83 to 0.88 and 0.90. However, increasing the concentration of the ionic liquid
further resulted in the formation of aggregates in the cationic polymer solution. Therefore,
NaCl was used to further optimize the separation conditions.
4.3.2 Effect of NaCl Concentration
Polymer deposition solutions composed of 0.5% (w/v) polymer and 0.00 M, 0.01
M, 0.05 M, and 0.10 M NaCl were used to study the influence of NaCl concentration on
the chiral separation of BNP. As shown in Figure 4.3, the retention times of the analyte
108
peaks slightly increased when NaCl was added to the solutions, probably due to an
increase in the thickness of the coating. In addition, an increase in resolution at higher
concentrations of NaCl was observed. The electroosmotic mobility decreased from 3.89 x
10-4 to 3.37 x 10-4 cm2V-1s-1 with increasing NaCl concentration. Based on these results,
0.1 M NaCl was used to further optimize the chiral separation of BNP.
4.3.3 Effect of Column Temperature
The influence of column temperature was also investigated. Figure 4.4 illustrates
the chiral separation of BNP at 35 °C, 25 °C and 15 °C. At a constant applied voltage, with
temperature decreasing, the retention times of R-(+)-BNP and S-(-)-BNP increased mainly
due to an increase in electrolyte viscosity. In addition, the electroosmotic mobility
decreased from 4.24 x 10-4 to 2.79 x 10-4 cm2V-1s-1 with decreasing temperature. In Figure
4.4, it is also shown that BNP was successfully resolved at a column temperature of 15 °C.
4.3.4 Effect of Bilayer Number
The effect of the number of bilayers on the resolution and analysis time of BNP
was also examined. It has been shown that an increase in the bilayer number resulted in an
enhanced film thickness [27, 36]. In this study, 2, 4, 8, and 12 bilayers were constructed in
the PEM coating. As shown in Figure 4.5, the analysis time, the resolution, and the
selectivity increased when more bilayers were constructed. However, two and four bilayers
did not demonstrate a change in selectivity, even though there was an increase in retention
time.
109
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.008
-0.006
-0.004
-0.002
0.000
(a) 0.01 M NaCl R-(+)-BNP S-(-)-BNP
17
O
O
PO
OH
αααα=1.022
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.010
-0.008
-0.006
-0.004
-0.002
0.000 (c) 0.01 M 1B-3MI-TFB
17
αααα=1.0235
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.004
-0.003
-0.002
-0.001
0.000
0.001
(b) 0.01 M 1E-3MI-HFP
17
αααα=1.023
Figure 4.2 Chiral separation of BNP. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV); pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); applied voltage, 30 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm. (a) 0.01 M NaCl; (b) 0.01 M 1E-3MI-HFP; (c) 0.01 M 1B-3MI-TFB.
110
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.010
-0.005
0.000
0.005
0.010
0.00 M NaCl
17
αααα=1.021
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.008
-0.006
-0.004
-0.002
0.000
0.01 M NaCl
17
αααα=1.022
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.008
-0.006
-0.004
-0.002
0.000
0.05 M NaCl
17
αααα=1.025
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.005
-0.004
-0.003
-0.002
-0.001
0.10 M NaCl
17
αααα=1.026
Figure 4.3 Effect of NaCl concentration on the chiral separation of BNP. Conditions: same as Figure 4.2, except NaCl concentration was varied.
111
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.004
-0.002
0.000
0.002 35 °C
17
αααα=1.019
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.005
-0.004
-0.003
-0.002
-0.001 25 °C
17
αααα=1.026
Minutes0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16
AU
-0.004
-0.002
0.000
0.002
15 °C
17
αααα=1.044
Figure 4.4 Effect of column temperature on the chiral separation of BNP. Conditions: same as Figure 4.2, except temperature was varied.
112
M inutes0 2 4 6 8 10 12 14 16 18 20 22 24 26
AU
-0.006
-0.005
-0.004
-0.003
2 Bilayers
αααα=1.026
M inute s0 2 4 6 8 10 12 14 16 18 20 22 24 26
AU
-0.002
-0.001
0.000
0.001
4 Bilayers
αααα=1.026
M inute s0 2 4 6 8 10 12 14 16 18 20 22 24 26
AU
-0.002
0.000
0.002
0.004
0.006
12 Bilayers
αααα=1.043
M inute s0 2 4 6 8 10 12 14 16 18 20 22 24 26
AU
-0.002
-0.001
0.000
0.001
8 Bilayers
αααα=1.040
Figure 4.5 Effect of bilayer number on the chiral separation of BNP. Conditions: same as Figure 4.2, except bilayer number was varied.
113
4.3.5 Reproducibilities
Reproducibility of the PEM coating is an important factor for the evaluation of
column performance. Reproducibilities were evaluated by using the relative standard
deviation (RSD) [37] values of the EOF and the R-(+)-BNP peak. In both cases, the run-to-
run RSD values were obtained from 10 consecutive electrophoresis runs. For each NaCl
concentration added to the polymer solutions (0.00 M, 0.01 M, 0.05 M, and 0.10 M), three
capillaries were coated. The RSD values of the EOF ranged from 0.13% to 0.49%, and the
RSD values of the R-(+)-BNP peak ranged from 0.24% to 0.99% (Table 4.1). Figure 4.6 is
an illustration of this excellent reproducibility, and it demonstrates 10 electropherograms
obtained from 10 replicate runs when 0.05 M NaCl was used.
The capillary-to-capillary reproducibilities were obtained from 30 runs (10
consecutive runs performed in 3 different capillaries for each NaCl concentration). It was
observed that all RSD values were below 0.5% (Table 4.2). In addition, 10 more replicate
analyses were performed in 6 consecutive days. For the first two days, the RSD values
were above 0.2% (Table 4.3). However, after the second day, all RSD values dropped
below 0.2%. Therefore, conditioning of the coated capillaries significantly improves the
reproducibility.
4.3.6 Chiral Separation of Analytes
As shown in Figure 4.7, four more analytes were able to be resolved with
resolution (Rs) values ranging from 1.07 to 1.55. BOH, which is a partially anionic analyte
at pH 10, was almost baseline separated with a resolution value of 1.24 (Figure 4.7a).
Temazepam is a neutral chiral compound in the benzodiazepine class of analytes. The
racemate of this analyte was successfully and easily resolved with the resolution value of
114
1.41 (Figure 4.7b). The barbiturates secobarbital and pentobarbital are partially anionic
compounds in a buffer solution of pH 7.2. The enantiomeric separation of both analytes
was performed using a three-bilayer PEM coating, 2.5% poly (L-SULV), and polymer
deposition solutions with 0.2 M NaCl. The chiral separations of both secobarbital (Figure
4.7c) and pentobarbital (Figure 4.7d) resulted in resolution values of 1.07 and 1.55,
respectively.
Table 4.1 Run-to-run reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration was varied.
RSD of R-BNP (%)RSD of EOF (%)
0.13
0.00 0.18
0.21
NaCl Concentration (M)
0.40
0.38
0.24
0.13
0.01 0.26
0.26
0.30
0.39
0.54
0.29
0.05 0.35
0.49
0.65
0.96
0.99
0.15
0.10 0.30
0.32
0.35
0.61
0.90
RSD of R-BNP (%)RSD of EOF (%)
0.13
0.00 0.18
0.21
NaCl Concentration (M)
0.40
0.38
0.24
0.13
0.01 0.26
0.26
0.30
0.39
0.54
0.29
0.05 0.35
0.49
0.65
0.96
0.99
0.15
0.10 0.30
0.32
0.35
0.61
0.90
115
Minutes
4.0 5.0 6.0 7.0 8.0 9.0 10.0 11.0
AU
0.00
0.02
0.04
0.06 0.05 M NaCl
12.0
Figure 4.6 Illustration of a run-to-run reproducibility for the chiral separation of BNP. Conditions: same as Figure 4.2, except NaCl concentration, 0.05 M.
116
Table 4.2 Capillary-to-capillary reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration was varied.
RSD of the EOF (%)
0.160.00
0.01
0.10
0.05
0.22
0.40
0.37
NaCl Concentration (M) RSD of the EOF (%)
0.160.00
0.01
0.10
0.05
0.22
0.40
0.37
NaCl Concentration (M)
Table 4.3 Reproducibilities of PEM capillary coating. Conditions: same as Figure 4.2, except NaCl concentration, 0.1 M.
RSD of the EOF (%)
0.30Day 1
Day 2
Day 4
Day 3
0.23
0.09
0.12
Day 6
Day 5 0.10
0.10
RSD of the EOF (%)
0.30Day 1
Day 2
Day 4
Day 3
0.23
0.09
0.12
Day 6
Day 5 0.10
0.10
117
M in u te s1 0 . 0 1 0 .5 1 1 .0 1 1 .5 1 2 .0 1 2 .5 1 3 .0 1 3 . 5 1 4 . 0 1 4 . 5 1 5 .0
A U
- 0 . 0 0 3 4
- 0 . 0 0 3 2
- 0 . 0 0 3 0
- 0 . 0 0 2 8
αααα=1.029
BO H
M in ute s5 .0 5 .2 5 .4 5 .6 5 .8 6 .0 6 .2 6 .4 6 .6 6 .8 7 .0 7 .2 7 .4
A U
0 .0 0 0
0 .0 0 2
0 .0 0 4
0 .0 0 6
0 .0 0 8
0 .0 1 0
Pentobarbital
αααα=1.184
M in u te s5 .1 5 .2 5 .3 5 .4 5 .5 5 . 6 5 . 7 5 . 8 5 .9 6 . 0 6 . 1 6 . 2
A U
0 .0 0 0
0 .0 0 1
0 .0 0 2
0 .0 0 3
0 .0 0 4
Secobarbital
αααα=1.105
M in u te s1 1 . 0 1 1 .2 1 1 .4 1 1 . 6 1 1 . 8 1 2 .0 1 2 .2 1 2 .4 1 2 .6 1 2 . 8 1 3 . 0 1 3 . 2 1 3 .4 1 3 .6 1 3 .8 1 4 .0 1 4 . 2 1 4 . 4 1 4 .6
A U
0 . 0 0 0 1
0 . 0 0 0 2
0 . 0 0 0 3
0 . 0 0 0 4
N
N
O H
O
Cl
H 3 C
Temazepam
αααα=1.032
(a)
(b)
(c)
(d)
Figure 4.7 (a) Chiral separation of BOH. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); detection, 214 nm. (b) Chiral separation of temazepam. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.9 psi for 7 s; background electrolyte, 25 mM Tris and 25 mM Na2B4O7 (pH 8.5); detection, 220 nm. (c) Chiral separation of secobarbital, and (d) Chiral separation of pentobarbital. Conditions for (c) and (d): 3 bilayers; 0.5% (w/v) PDADMAC and 2.5% (w/v) poly (L-SULV) with 0.2 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 300 mM H3BO3 and 30 mM Na2HPO4 (pH 7.2); detection, 214 nm. Other conditions are the same as Figure 4.2.
118
4.4 Conclusion
In this study, we were able to develop a simple PEM coating procedure for chiral
separations. The anionic polymer poly (L-SULV) proved to be a good chiral discriminator
for the separation of several analytes. The enantiomeric separations of BNP, BOH, and
temazepam were achieved using a two-bilayer PEM coating, 0.5% poly (L-SULV), and 0.1
M NaCl in the polymer deposition solutions. On the other hand, secobarbital and
pentobarbital were separated using a three-bilayer PEM coating, 2.5% poly (L-SULV), and
polymer deposition solutions with 0.2 M NaCl. In addition, the endurance of the PEM
coated capillary was more than 230 runs with RSD values of less than 1%. In all cases, the
run-to-run and capillary-to-capillary RSD values of EOF were less than 0.5%, and the run-
to-run RSD values of the R-(+)-BNP peak were less than 1%. It should be noted that the
chiral separation coating had to be modified from the procedure previously used for achiral
separations [18]. In our laboratory, further studies are ongoing to separate more chiral
compounds, and to better understand the structure of the coating.
4.5 References
1. Wang, J.; Warner, I. M. Anal. Chem. 1994, 66, 3773.
2. Jamali, F.; Mehvar, R.; Pasutto, F. M. J. Pharm. Sci. 1989, 78, 695.
3. Snopek, J.; Jelinek, I.; Smolkova-Keulemansova, E. J. Chromatogr. 1992, 609, 1.
4. Haddadian, F.; Billiot, E. J.; Shamsi, S. A.; Warner, I. M. J. Chromatogr. A 1999, 858, 219.
5. Shamsi, S. A.; Warner, I. M. Electrophoresis 1997, 18, 179.
6. Palmer, C. P.; Terabe, S. Anal. Chem. 1997, 69, 1852.
7. Wang, J.; Warner, I. M. J. Chromatogr. 1995, 711, 297.
119
8. Shamsi, S. A.; Valle, B. C.; Billiot, F.; Warner, I. M. Anal. Chem. 2003, 75, 379.
9. Pundlett, K. L.; Armstrong, D. W. Anal. Chem. 1996, 68, 3493.
10. Shamsi, S. A.; Macossay, J.; Warner, I. M. Anal. Chem. 1997, 69, 2980.
11. Billiot, F. H.; Billiot, E. J.; Warner, I. M. J. Chromatogr. A 2001, 922, 329.
12. Billiot, E.; Warner, I. M. Anal. Chem. 2000, 72, 1740.
13. Erim, F. B.; Cifuentes, A.; Poppe, H.; Kraak, J. C. J. Chromatogr. A 1995, 708, 356.
14. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 2254.
15. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 5272.
16. Graul, T. W.; Schlenoff, J. B. Anal. Chem. 1999, 71, 4007.
17. Rmaile, H. H.; Schlenoff, J. B. J. Am. Chem. Soc. 2003, 125, 6603.
18. Kapnissi, C. P.; Akbay, C.; Schlenoff, J. B.; Warner, I. M. Anal. Chem. 2002, 74,
2328.
19. Kamande, M. W.; Kapnissi, C. P.; Zhu, X.; Akbay, C.; Warner, I. M. Electrophoresis 2003, 24, 945.
20. Hayes, J. D.; Malik, A. Anal. Chem. 2001, 73, 987.
21. Decher, G.; Schmitt, J. Prog. Colloid Polym. Sci. 1992, 89, 160.
22. Decher, G. Science 1997, 277, 1232.
23. Schlenoff, J. B.; Ly, H.; Li, M. J. Am. Chem. Soc. 1998, 120, 7626.
24. Sui, Z.; Salloum, D.; Schlenoff, J. B. Langmuir 2003, 19, 2491.
25. Schlenoff, J. B.; Dubas, S. T.; Farhat, T. Langmuir 2000, 16, 9968.
26. Caruso, F.; Lichtenfeld, H.; Donath, E.; Möhwald, H. Macromolecules 1999, 32,
2317.
27. Laurent, D.; Schlenoff, J. B. Langmuir 1997, 13, 1552.
28. Smith, R. N.; Reven, L.; Barrett, C. J. Macromolecules 2003, 36, 1876.
120
29. Rmaile, H. H.; Schlenoff, J. B. Langmuir 2002, 18, 8263.
30. Liu, Y.; Fanguy, J. C.; Bledsoe, J. M.; Henry, C. S. Anal. Chem. 2000, 72, 5939.
31. Dubas, S. T.; Schlenoff, J. B. Macromolecules 1999, 32, 8153.
32. Lösche, M.; Schmitt, J.; Decher, G.; Bouwman, W. G.; Kjaer, K. Macromolecules 1998, 31, 8893.
33. Armstrong, D. W.; He, L.; Liu, Y. Anal. Chem. 1999, 71, 3873.
34. Dupont, J.; De Souza, R. F.; Suarez, P. A. Z. Chem. Rev. 2002, 102, 3667.
35. Yanes, E. G.; Gratz, S. R.; Baldwin, M. J.; Robison, S. E.; Stalcup, A. M. Anal.
Chem. 2001, 73, 3838.
36. Schlenoff, J. B.; Dubas, S. T. Macromolecules 2001, 34, 592.
37. Skoog, D. A.; Holler, J. F.; Nieman, T. A. In Principles of Instrumental Analysis, 5th ed.; Sherman, M., Bortel, J., Messina, F., Eds.; Harcourt Brace College Publishers: Orlando, FL, 1998.
121
CHAPTER 5.
INVESTIGATION OF THE STABILITY OF POLYELECTROLYTE MULTILAYER COATINGS IN OPEN-TUBULAR CAPILLARY
ELECTROCHROMATOGRAPHY USING LASER SCANNING CONFOCAL MICROSCOPY
5.1 Introduction
In recent years, layer-by-layer deposition process of polyelectrolytes on hydrophilic
surfaces via electrostatic interactions has attracted considerable attention. This method was
first developed by Decher et al. [1, 2]. In this approach, a substrate is alternately exposed
to solutions of positively and negatively charged polyelectrolytes. The alternate adsorption
of these oppositely charged polyelectrolytes produces fairly uniform thin films that have
been used in the areas of light-emitting devices, sensors, coatings, nonlinear optics,
catalysis, patterning, diagnostics, bioadhesion, and drug delivery [3-6].
The layer-by-layer method has also been involved in analytical separations [7-11].
This coating involves a polyelectrolyte multilayer (PEM), constructed in a fused-silica
capillary by alternating rinses of cationic and anionic polyelectrolytes [12-22]. The first
layer of the cationic polymer adds to the negatively charged surface of the capillary. The
construction of the cationic layer reverses the charge on the surface and primes the film for
the addition of the next layer of anionic polymer. These layer-by-layer coatings have
shown strong stability and high resistance to charge and deterioration during use [7, 8, 23,
24].
Katayama et al. [23, 24] established a stable modification of the inner walls of the
capillary by use of the successive multiple ionic-polymer layer (SMIL) coating. Their first
study focused on anion-modified capillaries that were achieved by first attaching the
122
cationic polymer to the inner walls of the capillaries and then, the anionic polymer to the
cationic polymer layer [23]. The endurance of the coated capillary was more than 100 runs,
and this is due to the multiple attachment of the anionic polymer. The coating was also
tolerant to some organic and alkaline solvents. However, the coated capillary was unstable
after 0.1 M HCl and CH3CN rinsing, and the coating was detached after 1 M NaOH rinse.
In their second study, these authors developed a cation-modified capillary by attaching the
cationic polymer to the anionic polymer layer [24]. This coated capillary endured 600
replicate runs, and showed strong stability against 1 M NaOH and 0.1 M HCl with
degradation ratios of less than 2%. Degradation ratios are changes in electroosmotic flow
(EOF).
Graul and Schlenoff [7] coated fused-silica capillaries with thin films of charged
polymers by use of the PEM coating procedure. They demonstrated that these columns
were stable to extremes of pH and ionic strength, and to dehydration/rehydration. For the
stability studies against exposure to extreme pH values and ionic strengths, the 6.5-layer
pair-coated capillary was flushed twice with 0.01 M (pH 12) for 10 min, and twice with
0.01 M HCl (pH 2) for 10 min.
In the study reported in this chapter, we describe an approach that uses open-
tubular capillary electrochromatography (OT-CEC) and laser scanning confocal
microscopy (LSCM) to further investigate the stability of the PEM coating approach that
was previously described. In this approach, the polymeric surfactant poly (sodium N-
undecanoyl-L-leucylvalinate), poly (L-SULV), was used in the construction of the coating.
Using OT-CEC, both electroosmotic mobility and selectivity changes were measured after
flushing the capillaries with 0.1 M and 1.0 M NaOH. In addition, a correlation between
123
flushing time and change in electroosmotic mobility and selectivity was investigated. The
structural changes of these coatings were also monitored and imaged after exposure to 1.0
M NaOH with LSCM. LSCM has proven to be a useful and valuable tool for
characterization of packed columns, coatings, tablets, capsules, living cells, and other
surfaces [25-39].
5.2 Experimental
5.2.1 Apparatus and Conditions
(i) Capillary Electrochromatography
All experiments were performed with a Beckman P/ACE MDQ capillary
electrophoresis system using UV detection (Fullerton, CA). Fused-silica capillaries of 57
cm (50 cm effective length) x 50 µm i.d., were purchased from Polymicro Technologies
(Phoenix, AZ). The capillaries were thermostated at 25 °C by use of liquid coolant. The
applied voltage was 30 kV, and the samples were injected by pressure at 0.5 psi (1 psi =
6894.76 Pa) for 3 s. UV detection was done at 214 nm.
(ii) Laser Scanning Confocal Microscopy
An inverted confocal microscope (Axiovert 100, LSM 510 scan module, Carl Zeiss
Inc., Thornwood, NY) was used to image the PEM-coated capillaries. The setup for
imaging has been described elsewhere [25]. Briefly, a 4-cm coated capillary was placed
between a microscope slide and a coverslip supported by two spacers consisting of two
short pieces of fused-silica capillary. The window of the coated capillary was immersed in
a refractive index matching fluid. Refractive index matching is important for obtaining
minimally distorted images by reduction of refraction and scattering from the capillary
walls. This arrangement is then placed on the microscope stage.
124
The imaging system was equipped with a helium-neon laser (Uniphase, 1674P) that
provided excitation at 543 nm. All confocal images were obtained with a 20x (NA 0.5) dry
objective. Band-pass or long-pass filters were used for selection of the spectral range of
fluorescence emission. Fluorescence was directed through an adjustable pinhole to a
photomultiplier tube.
5.2.2 Reagents and Chemicals
The analyte 1,1’-binaphthyl-2,2’-dihydrogenphosphate (BNP), and the buffer
tris(hydroxymethyl)aminomethane (Tris) were purchased from Sigma Chemical Co. (St.
Louis, MO). Sodium borate (Na2B4O7), sodium chloride (NaCl), and sodium hydroxide
(NaOH) were obtained from Fisher Scientific (Fair Lawn, NJ). Rhodamine 6G (R6G) and
PDADMAC (Mw = 200,000-350,000) were obtained from Aldrich (Milwaukee, WI).
Refractive index matching oil (n=1.51) was purchased from Carl Zeiss Microimaging, Inc.
(Thornwood, NY). The dipeptide L-leucylvalinate, undecylenic acid, and N-
hydroxysuccinimide were also obtained from Sigma.
5.2.3 Sample and Background Electrolyte Preparation
The analyte BNP was dissolved in 50:50 methanol/water. Its final concentration
was 0.1 mg/ml. The background electrolyte (BGE) consisted of 100 mM Tris and 10 mM
Na2B4O7. The pH was adjusted to 10 by use of 1.0 M NaOH. The solution was filtered
using 0.45 µm polypropylene nylon filters and sonicated for 15 min before use. The
fluorescent dye R6G was dissolved in deionized water at concentrations of 10 µM and 100
µM.
125
5.2.4 Synthesis of Polymeric Surfactant
The monomeric surfactant of sodium N-undecenoyl-L-leucylvalinate was
synthesized in our laboratory using the N-hydroxysuccinimide ester of undecylenic acid
according to a procedure previously described by Wang and Warner [40]. A 100 mM
sodium salt solution of the monomer was then polymerized by use of 60Co-γ radiation to
form the polymeric surfactant poly (L-SULV). The structure of poly (L-SULV) is shown in
Figure 4.1 (Chapter 4).
5.2.5 Procedure for Polyelectrolyte Multilayer Coating
The procedure for construction of the PEM coating has been described in Chapters
3 and 4 [8]. The cationic and anionic polymer deposition solutions were alternately flushed
through the capillary by use of the rinse function on the Beckman CE system. Both
solutions contained 0.5% (w/v) polymer in 0.1 M aqueous NaCl solution. First, the fused-
silica capillary was conditioned with water for 5 min, 1.0 M NaOH for 60 min, and again
with water for 15 min. Then, the first layer of the polymer PDADMAC was constructed by
rinsing the polymer solution through the capillary for 5 min followed by a water rinse for
another 5 min. All other polymer depositions were performed by flushing the capillary
with the polymer solutions for 5 min followed by 5-min water rinses. The multilayer
coatings used for the OT-CEC and LSCM studies consisted of two and twenty layer pairs.
For the LSCM experiments, after each coating was constructed, the capillary was flushed
with 100 µM R6G for 10 min in order to incorporate the fluorescent probe into the
polymer coating. For the chromatographic experiments, after the construction of the
coating, the capillary was conditioned with the BGE until a stable current was achieved.
126
5.3 Results and Discussion
5.3.1 Open-Tubular Capillary Electrochromatography
Experiments performed in OT-CEC studied the stability of capillary surface, the
capillary recovery, and the coating regeneration. These parameters were evaluated by
computing the electroosmotic mobility, EOFµ , and the selectivity factor, α .
As stated in Chapter 2, the electroosmotic mobility is described by the following
equation:
0VtLL td
eo =µ (5.1)
where dL is the distance from injector to detector; tL is the total length of the capillary; 0t
is the migration time of the EOF marker; and V is the applied voltage. Methanol was used
as the EOF marker in this study since it is not retained by the PEM coating.
Selectivity is another important parameter that was used to study the stability and
regeneration of the coating after exposure to NaOH. The selectivity factor of a capillary
column for two species A and B, as already discussed, is defined as:
( )( ) 0
0
tttt
Ar
Br
−−
=α (5.2)
where ( )Brt and ( )Art are the retention times of B and A, respectively. In this study, B and
A are the enantiomers S-(-)-BNP and R-(+)-BNP, respectively.
5.3.1.1 Coating Stability
To study the stability of the capillary surface, two bilayers were constructed. First,
the EOF was measured before the coating was deposited on the capillary walls of the
fused-silica capillary. The measured electroosmotic mobility was 5.04 x 10-4 cm2V-1s-1.
127
The EOF was again measured after the two bilayers were constructed in the PEM coating.
In this case, the electroosmotic mobility decreased to 3.31 x 10-4 cm2V-1s-1. Then, 0.1 M
and 1.0 M NaOH solutions were flushed through the PEM-coated capillary, and the
magnitude of the EOF was measured. Figure 5.1 demonstrates the dependence of
electroosmotic mobility on the NaOH flushing time. In addition, this figure illustrates
graphically how NaOH can increase the value of electroosmotic mobility, as compared to
the 2-bilayer capillary that is not treated with the strong base. As shown, 0.1 M NaOH did
not have a major impact on electroosmotic mobility, as compared to 1.0 M NaOH, even
after flushing the coated capillary for 100 min. A 5-min 1.0 M NaOH flushing time slightly
increased the value of electroosmotic mobility to 3.35 x 10-4 cm2V-1s-1. However, fifteen
more minutes demonstrated a significant increase that was still observed upon continuous
rinse with 1.0 M NaOH. A total of 230 min resulted in an electroosmotic mobility of 5.03
x 10-4 cm2V-1s-1. In addition, a total of 350 min gave the same EOF value obtained before
deposition of the coating (5.04 x 10-4 cm2V-1s-1). This clearly demonstrates that the coating
was completely desorbed from the capillary column. It should be noted that each data point
in this graph and the graphs that follow is the average value obtained from running each
experiment multiple times. Standard deviation values were also calculated for each data
point. However, these values are very small, and not perceptible on the graphs.
Figure 5.2 demonstrates the dependence of selectivity on the NaOH flushing time.
Before polymer deposition, the selectivity was 1.0 since no separation between the two
enantiomers of BNP was observed. The construction of two bilayers gave a selectivity
value of 1.026, which was relatively stable during the coating’s exposure to 0.1 M NaOH.
A slight decrease was observed when 1.0 M NaOH solution was flushed through the
128
coated capillary. A total flush time of 170 min with 1.0 M NaOH resulted in a single BNP
peak. This observation also confirmed that the coating was removed after a long exposure
to NaOH.
3
3.5
4
4.5
5
5.5
0 20 40 60 80 100 120Time (minutes)
3
3.5
4
4.5
5
5.5
0 50 100 150 200 250 300 350 400
EOF
(cm
2 V-1
s-1 x
10-4
)
time (minutes)
0.1 M NaOH 1.0 M NaOH
Figure 5.1 Dependence of electroosmotic mobility on the NaOH flushing time. Conditions: 2 bilayers; 0.5% (w/v) PDADMAC and 0.5% (w/v) poly (L-SULV) with 0.1 M NaCl; pressure injection, 0.5 psi for 3 s; background electrolyte, 100 mM Tris and 10 mM Na2B4O7 (pH 10.0); applied voltage, 30 kV; temperature, 25 °C; capillary, 57 cm (50 cm effective length) x 50 µm i.d.; detection, 214 nm.
The stability was further evaluated by calculating the change in EOF, also termed
“degradation ratio” [23, 24]. This ratio was calculated by use of the following equation:
Degradation ratio = 1001
21 ×−
EOFEOFEOF
(5.3)
129
where 1EOF and 2EOF were measured before and after flushing the capillary with the
NaOH solution, respectively. In this case, 1EOF is as mentioned above, i.e. 3.31 x 10-4
cm2V-1s-1. These degradation ratio values are reported in Table 5.1. The PEM-coated
capillary was relatively stable after rinsing with 0.1 M NaOH for 110 min, followed by a
rinse with 1.0 M NaOH for 5 min. The degradation ratios obtained until the last exposure
(1.0 M NaOH for 5 min) were all below 2%. However, after additional 1.0 M NaOH rinse,
the degradation ratios were above 2%. The coating was finally detached from the capillary
wall after long exposure to 1.0 M NaOH as evidenced by α and the EOF value.
0.99
1
1.01
1.02
1.03
1.04
1.05
0 20 40 60 80 100 120Time (minutes)
0.99
1
1.01
1.02
1.03
1.04
1.05
0 50 100 150 200 250 300 350 400
Sele
ctiv
ity
time (minutes)
0.1 M NaOH 1.0 M NaOH
Figure 5.2 Dependence of selectivity on the NaOH flushing time. Conditions: same as Figure 5.1.
130
Table 5.1 Stability of PEM capillary coating. Conditions: same as Figure 5.1.
Degradation Ratio(%)
EOF2 Average (min)(x 10-4 cm2 V-1s-1)
5.0
Total Flushing Time(min)
0.1 M NaOH
20.0
50.0
110.0
1.0 M NaOH
5.0
20.0
50.0
110.0
170.0
230.0
350.0
3.31
3.31
3.31
3.26
0.00
0.00
0.00
1.51
3.35
3.65
3.86
4.39
4.45
5.03
5.04
1.21
10.27
16.61
32.63
34.44
51.96
52.27
5.3.1.2 Capillary Recovery and Coating Regeneration
When the coating was completely detached, and a selectivity value of 1.0 was
obtained, a 2-bilayer coating was reconstructed in the same fused-silica capillary. Figure
131
5.3 graphically demonstrates how electroosmotic mobility gradually changes after again
flushing the coated capillary with NaOH solutions. The trend observed in this figure is the
same as obtained when the first 2-bilayer coating was constructed. The EOF after the last
coating deposition was 3.32 x 10-4 cm2V-1s-1, as compared to the value of 3.31 x 10-4 cm2V-
1s-1 that was measured after the first coating. In addition, a similar trend was observed with
the selectivity values, as shown in Figure 5.4.
3
3.5
4
4.5
5
5.5
0 20 40 60 80 100 120Time (minutes)
3
3.5
4
4.5
5
5.5
0 50 100 150 200 250 300 350 400
0.1 M NaOH 1.0 M NaOH
EOF
(cm
2 V-1
s-1 x
10-4
)
time (minutes)
Figure 5.3 Coating Regeneration - Dependence of electroosmotic mobility on the NaOH flushing time. Conditions: same as Figure 5.1.
132
0.99
1
1.01
1.02
1.03
1.04
1.05
0 20 40 60 80 100 120Time (minutes)
0.99
1
1.01
1.02
1.03
1.04
1.05
0 50 100 150 200 250 300 350 400
Sele
ctiv
ity
time (minutes)
0.1 M NaOH 1.0 M NaOH
Figure 5.4 Dependence of selectivity on the NaOH flushing time. Conditions: same as Figure 5.1.
Another important consideration for this study is the amount of time that is needed
to completely detach the coating for capillary recovery. This allows us regeneration of the
coating by use of this simple PEM coating procedure. For purpose of this study, two and
twenty bilayers were constructed. Both electroosmotic mobility and selectivity
measurements illustrated that a 2-bilayer and a 20-bilayer PEM coating (data not shown)
could be completely detached from the capillary walls after approximately 3.5 and 9.5
hours, respectively of continuous exposure to 1.0 M NaOH.
133
5.3.2 Laser Scanning Confocal Microscopy
Using the results from the above study, one can conclude that the PEM coating
constructed using polymeric surfactants as anionic polymers affects separations [8-10, 44].
However, we desired an imaging technique that would allow visualization of the PEM-
coated capillaries for characterization of the surface morphology. In this part of our study,
LSCM was used as an imaging technique to examine the coating. LSCM was also used to
image the structural changes of the coating after its exposure to NaOH. For the
experiments reported here, R6G proved to be a suitable fluorescent dye to examine the
coating since this positively charged probe prefers the environment of the coated phase.
Figure 5.5 is a comparison of a 20-bilayer capillary (left) and a fused-silica
capillary (right) side by side in a single image. This comparison is used to visually
demonstrate the difference between a coated and bare capillary. In both sections of the
capillaries, 10 µM R6G in water was flushed, and then, air was injected in order to wash
the excess solution out and dry the capillary. This figure illustrates z-section images that
were collected at different depths of focus of the LSCM. The pinhole size of the LSCM
was adjusted such that the optical slice was 3.3 µm. From Figure 5.5a to Figure 5.5f, the z-
sectioning goes deeper into the PEM-coated capillary. The focal plane in Figure 5.5a is at
the outer edge of the capillary wall. Figure 5.5f slices through the outer wall of the
capillary on the opposite side. Figures 5.5c and 5.5d slice though regions of the capillary
near the center plane. As shown in these images, the walls of the 20-bilayer capillary are
very bright as compared to the fused-silica capillary. Since the R6G should be retained to a
greater extent in the coated phase than on the wall of the fused-silica capillary, this
difference confirms the existence of a coating on the capillary wall.
134
(a) (b)
(c) (d)
(e) (f)
20-bilayer capillary Fused-silica capillary
Figure 5.5 The z-section images of a 20-bilayer capillary (left) and a fused-silica capillary (right). Optical slice thickness: 3.3 µm. z-Step size 1.65 µm. Every fourth image is shown. All images are 204 x 512 pixels (460.6 µm x 115.1 µm).
135
The same 20-bilayer capillary was exposed to 1.0 M NaOH. It was then flushed
again with 10 µM R6G and injected with air. Using the confocal microscope, we observed
a change in the structure of the coating, i.e. large portions of the polymer coating have
been removed from the wall. This observation is clearly illustrated in Figure 5.6. The
images of this figure represent again z-section images that were taken as the excitation
beam traversed deeper into the coated capillary.
(a) (b) (c)
(d) (e) (f)
Figure 5.6 The z-section images of a portion of the same 20-bilayer capillary, as in Figure 5.5. Exposure to 1.0 M NaOH. Optical slice thickness: 3.3 µm. z-Step size 1.65 µm. Every third image is shown. All images are 512 x 512 pixels (65.8 µm x 65.8 µm).
136
Additional LSCM images of larger portions of the capillary at different locations
demonstrate similar discontinuities in the coating (Figure 5.7). The area depicted in Figure
5.7a includes the discontinuity and a large region where the coating on the walls of the
capillary does not appear to be damaged to the same extent. Figure 5.7b includes several
discontinuities along the wall of the capillary where the coating has been greatly affected.
(a) (b)
Figure 5.7 LSCM images of different regions within the same 20-bilayer capillary, as in Figure 5.6. Optical slice thickness: 3.3 µm. Both images are 512 x 2048 pixels (115.1 µm x 460.6 µm).
137
When the same 20-bilayer capillary was exposed to additional 1.0 M NaOH, the
majority of the coating appears to be removed, leaving only several bright islands on the
wall (Figure 5.8). In addition, this last image demonstrates that the coating is removed after
long exposure to 1.0 M NaOH. The same procedure was followed with the 2-bilayer
capillary (data not shown).
Figure 5.8 LSCM image of the same 20-bilayer capillary, as in Figure 5.7. Longer exposure to 1.0 M NaOH. Optical slice thickness: 3.3 µm. The image is 512 x 820 pixels (115.1 µm x 184.4 µm).
5.4 Conclusion
The stability of PEM coatings was further investigated in this study after exposure
to NaOH solutions. The OT-CEC study, which allowed measurements of electroosmotic
mobility and selectivity, suggests that the PEM coating is relatively stable against 0.1 M
NaOH. In addition, both OT-CEC and LSCM demonstrated the ability to recover the
138
capillaries by flushing them with 1.0 M NaOH. We were also able to visualize the coating
within the capillary, and to study its structural changes by using LSCM. This helped us
prove that the coating exists, allowed us to see the effects of NaOH on the coating, and
demonstrated that the coating can be removed after long exposure to 1.0 M NaOH.
Although LSCM provided new information, it did not provide us with all the details
regarding the structure of the coating.
5.5 References
1. Decher, G.; Schmitt, J. Prog. Colloid Polym. Sci. 1992, 89, 160.
2. Decher, G. Science 1997, 277, 1232.
3. Schlenoff, J. B.; Dubas, S. T.; Farhat, T. Langmuir 2000, 16, 9968.
4. Smith, R. N.; Reven, L.; Barrett, C. J. Macromolecules 2003, 36, 1876.
5. Schlenoff, J. B.; Ly, H.; Li, M. J. Am. Chem. Soc. 1998, 120, 7626.
6. Farhat, T. R.; Schlenoff, J. B. Langmuir 2001, 17, 1184.
7. Graul, T. W.; Schlenoff, J. B. Anal. Chem. 1999, 71, 4007.
8. Kapnissi, C. P.; Akbay, C.; Schlenoff, J. B.; Warner, I. M. Anal. Chem. 2002, 74, 2328.
9. Kamande, M. W.; Kapnissi, C. P.; Zhu, X.; Akbay, C.; Warner, I. M.
Electrophoresis 2003, 24, 945.
10. Kapnissi, C. P.; Valle, B. C.; Warner, I. M. Anal. Chem. 2003, 75, 6097.
11. Rmaile, H. H.; Schlenoff, J. B. J. Am. Chem. Soc. 2003, 125, 6602.
12. Müller, M. Biomacromolecules 2001, 2, 262.
13. Laurent, D.; Schlenoff, J. B. Langmuir 1997, 13, 1552.
14. Schlenoff, J. B.; Dubas, S. T. Macromolecules 2001, 34, 592.
139
15. Caruso, F.; Lichtenfeld, H.; Donath, E.; Möhwald, H. Macromolecules 1999, 32, 2317.
16. Rmaile, H. H.; Schlenoff, J. B. Langmuir 2002, 18, 8263.
17. Castelnovo, M.; Joanny, J.-F. Langmuir 2000, 16, 7524.
18. Dubas, S. T.; Schlenoff, J. B. Macromolecules 2001, 34, 3736.
19. Dubas, S. T.;Schlenoff, J. B. Macromolecules 1999, 32, 8153.
20. Hayes, J. D.; Malik, A. Anal. Chem. 2001, 73, 987.
21. Farhat, T.; Yassin, G.; Dubas, S. T.; Schlenoff, J. B. Langmuir 1999, 15, 6621.
22. Sui, Z.; Salloum, D.; Schlenoff, J. B. Langmuir 2003, 19, 2491.
23. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 2254.
24. Katayama, H.; Ishihama, Y.; Asakawa, N. Anal. Chem. 1998, 70, 5272.
25. Lowry, M.; He, Y.; Geng, L. Anal. Chem. 2002, 74, 1811.
26. Zhu, H.; Ji, J.; Tan, Q.; Barbosa, M. A.; Shen, J. Biomacromolecules 2003, 4, 378.
27. Moya, S.; Donath, E.; Sukhorukov, G. B.; Auch, M.; Bäumler, H.; Lichtenfeld, H.;
Möhwald, H. Macromolecules 2000, 33, 4538.
28. Guo, H. X.; Heinämäki, J.; Yliruusi, J. International J. Pharmaceutics 1999, 186, 99.
29. Silvano, D.; Krol, S.; Diaspro, A.; Cavalleri, O.; Gliozzi, A. Microscopy Research
and Technique 2002, 59, 536.
30. Georgieva, R.; Moya, S.; Leporatti, S.; Neu, B.; Bäumler, H.; Reichle, C.; Donath, E.; Möhwald, H. Langmuir 2000, 16, 7075.
31. Voigt, A.; Lichtenfeld, H.; Sukhorukov, G. B.; Zastrow, H.; Donath, E.; Bäumler,
H.; Möhwald, H. Ind. Eng. Chem. Res. 1999, 38, 4037.
32. Lamprecht, A.; Schäfer, U. F.; Lehr, C.-M. European J. Pharmaceutics and Biopharmaceutics 2000, 49, 1.
33. Stricker, S. A.; Whitaker, M. Microscopy Research and Technique 1999, 46, 356.
140
34. Gao, C.; Leporatti, S.; Donath, E.; Möhwald, H. J. Phys. Chem. B 2000, 104, 7144.
35. Buttino, I.; Ianora, A.; Carotenuto, Y.; Zupo, V.; Miralto, A. Microscopy Research and Technique 2003, 60, 458.
36. Khopade, A. J.; Caruso, F. Langmuir 2003, 19, 6219.
37. Astrakharchik-Farrimond, E.; Shekunov, B. Y.; Sawyer, N. B. E.; Morgan, S. P.;
Somekh, M. G.; See, C. W. Particle and Particle Systems Characterization 2003, 20, 104.
38. Missert, N.; Copeland, R. G.; Barbour, J. C.; Mikkalson, J. E. Electrochemical
Society Proceedings 2001, 22, 687.
39. Caponetti, G.; Hrkach J. S.; Kriwet, B.; Poh, M.; Lotan, N.; Colombo, P.; Langer, R. J. Pharm. Sci. 1999, 88, 136.
40. Wang, J.; Warner, I. M. J. Chromatogr. 1995, 711, 297.
41. Chiari, M.; Nesi, M.; Righetti, P. G. In Capillary Electrophoresis in Analytical
Biotechnology; Righetti, P. G., Ed.; CRC Press, Inc.: Boca Raton, FL, 1996.
42. Chankvetadze, B. In Capillary Electrophoresis in Chiral Analysis; John Wiley & Sons Ltd: London, UK, 1997.
43. Skoog, D. A.; Holler, J. F.; Nieman, T. A. In Principles of Instrumental Analysis,
5th ed.; Sherman, M., Bortel, J., Messina, F., Eds.; Harcourt Brace College Publishers: Orlando, FL, 1998.
44. Zhu, X.; Kamande, M. W.; Thiam, S.; Kapnissi, C. P.; Mwongela, S. M.; Warner, I.
M. Electrophoresis, 2004, 25, 562.
141
CHAPTER 6.
CONCLUSIONS AND FUTURE STUDIES
In this work, two different modes of capillary electrochromatography (CEC) were
used for the achiral and chiral separations of various classes of analytes. In Chapter 2 of
this dissertation, the packed mode of CEC is described. A 40-cm packed bed of Reliasil 3
µm C18 stationary phase was able to separate seven benzodiazepines. The separation
conditions were optimized by varying the mobile phase, the amount of organic modifier,
the buffer concentration, the applied voltage, and the column temperature. A mobile phase
composition of Tris.HCl (pH 8)-acetonitrile (60:40), an electrolyte concentration of 30
mM, and a temperature of 15 °C with an applied voltage of 20 kV proved to be optimum.
In addition, the packed capillary electrochromatographic method developed here was
applied to the characterization of oxazepam in a standard urine sample that contained
various concentrations of other drugs, such as acetaminophen, amphetamines, imipramine,
morphine, cocaine, etc.
Some preliminary data were also obtained for the separation of a mixture of six β-
blockers using packed-CEC with octadecyl silica stationary phase and co-polymerized
micelle SUS/SUG (sodium undecylenic sulfate/sodium N-undecanoyl-L-glycinate)
“pseudostationary phase.” β-blockers are clinically important drugs, used for the treatment
of several disorders such as angina pectoris, cardiac arrhythmias, and hypertension. They
have also been used as doping agents for athletes in order to improve their athletic
performance in cases where high psychological pressure may cause the heart to race. The
structures of the β-blockers used in this study are shown in Figure 6.1. Co-polymerized
micelles are produced by irradiating a micellar solution of surfactants with 60Co
142
irradiation. Their main advantage is that they can be used in a wider pH range than the
conventional polymeric surfactants, since they are soluble at and below pH 6.8. The
studies that have been performed until now showed that the presence of the co-
polymerized micelle SUS/SUG improved the resolution of the analytes. The resolution was
also proportional to the concentration of the co-polymerized micelle and the percentage of
SUS in the co-polymerized micelle. The studies on the separation of β-blockers will be
continued, and when optimum conditions are established, the method will be applied to a
real sample (urine or blood).
1. Sotalol 2. Atenolol
4. Labetalol
5. Propranolol 6. Acebutolol
3. Metoprolol
CHCH2NHCH(CH3)2CH3SO2NH
OH
OCH2CHCH2NHCH(CH3)2H2NCCH2
O OH
OCH2CHCH2NHCH(CH3)2CH3OCH2CH2
OH
HO
H2NC
CHCH2NHCHCH2CH2
O
CH3
OH
OCH2CHCH2NHCH(CH3)2
OH
CCH3
O
OCH2CHCH2NHCH(CH3)2
OH
CH3CH2CH2CNH
O
Figure 6.1 Structures of the six β-blocker analytes.
143
Chapter 3 is an examination of the open-tubular mode of CEC, which is an
alternative approach to conventional CEC. The primary advantage of open-tubular CEC
(OT-CEC) is the elimination of problems associated with frits and silica particles in
conventional CEC. In this approach, fused silica capillaries coated with thin films of
physically adsorbed charged polymers were developed by use of a polyelectrolyte
multilayer (PEM) coating procedure. The PEM coating was constructed in situ by
alternating rinses with positively and negatively charged polymers, where the negatively
charged polymer was a polymeric surfactant. In this study, poly
(diallyldimethylammonium chloride), PDADMAC, was used as the cationic polymer and
poly (sodium N-undecanoyl-L-glycinate), poly (L-SUG), was used as the anionic polymer
for PEM coating. The performance of the modified capillaries as a separation medium was
evaluated by use of seven benzodiazepines as analytes. The run-to-run, day-to-day, week-
to-week and capillary-to-capillary reproducibilities of electroosmotic flow (EOF) were
very good with relative standard deviation (RSD) values of less than 1% in all cases. In
addition, the chromatographic performance of the monomeric form of the polymeric
surfactant was compared for the separation of these analytes. The anionic surfactant used
for PEM coating construction in this experiment was the monomeric (nonpolymerized)
surfactant at the concentration of 0.5% (w/v) (19 mM). Almost no separation was noted,
even though the monomeric surfactant concentration was above the normal critical
micellar concentration of the nonpolymerized surfactant (7 mM). Therefore, the molecular
micelle allows better discrimination of the hydrophobic analytes than the conventional
micelle. Furthermore, the PEM-coated capillary was remarkably robust with more than 200
144
runs accomplished in this study. Strong stability against extreme pH values (pH 11.0 and
pH 3.0) was also observed.
In Chapter 4, the fused-silica capillaries were again modified using the PEM
coating procedure. The quaternary ammonium salt PDADMAC was used as the cationic
polymer, and the polymeric surfactant poly (sodium N-undecanoyl-L-leucylvalinate), poly
(L-SULV), was used as the anionic polymer. In this study, the PEM coating was applied to
investigate the chiral separations of 1,1’-binaphthyl-2,2’-dihydrogenphosphate (BNP),
1,1’-bi-2-naphthol (BOH), secobarbital, pentobarbital and temazepam. However, the PEM
coating procedure used in the achiral studies needed to be modified in order to achieve
chiral separations. Optimal conditions were established by varying the additive (sodium
chloride, 1-ethyl-3-methyl-1H-imidazolium hexafluorophosphate (1E-3MI-HFP), 1-butyl-
3-methylimidazolium tetrafluoroborate (1B-3MI-TFB)) in the polymer deposition
solutions, the salt concentration, the column temperature, and the bilayer number. The
ionic liquids 1E-3MI-HFP and 1B-3MI-TFB were explored for possible enhanced
separations. The addition of either ionic liquid to the polymer deposition solutions
increased the resolution of BNP from 0.83 to 0.88 and 0.90. The effects that ionic liquids
have on separations are not very well understood. Therefore, fluorescence studies should
be performed to better understand the mechanisms that take place in separations when
ionic liquids are used as additives. Reproducibilities were also evaluated by using the RSD
values of the EOF and the first peak (R-(+)-BNP). In all cases, the run-to-run and
capillary-to-capillary RSD values of EOF were less than 0.5%, and the run-to-run RSD
values of the R-(+)-BNP peak were less than 1%. In addition, more than 230 runs were
performed in the PEM coated capillary.
145
The studies described above for both chiral and achiral separations have shown that
PEM-coated capillaries have excellent reproducibilities, remarkable endurance, and strong
stabilities against extreme pH values when used in open tubular capillary
electrochromatography (OT-CEC). In Chapter 5, the stability of the capillary surface was
further investigated after exposure to 0.1 M and 1.0 M NaOH. The multilayer coatings
used for the studies reported in this chapter consisted of two and twenty layer pairs. A
layer pair, i.e. a bilayer, is one layer of a cationic polymer and one layer of an anionic
polymer. PDADMAC was used as the cationic polymer, and the polymeric surfactant poly
(L-SULV) was used as the anionic polymer. The structural changes of these coatings were
monitored using laser scanning confocal microscopy (LSCM) after flushing the capillaries
with NaOH. This technique also allowed a study of the uniformities and discontinuities of
the coatings. The observed structures were discussed in terms of separations using OT-
CEC. In addition, the electropherograms obtained from the chiral separation of BNP in
OT-CEC showed a decrease in selectivity and an increase in electroosmotic mobility after
long exposure to NaOH. To study the stability of the capillary surface, two bilayers were
constructed. The stability was evaluated by computing the change in EOF (degradation
ratio). The degradation ratios obtained after rinsing the PEM-coated capillary with 0.1 M
NaOH for 110 min, and with 1.0 M NaOH for 5 min were all below 2%. However, after
additional 1.0 M NaOH rinse, the degradation ratios were above 2%. Finally, the ability to
recover the capillaries by direct exposure to NaOH was demonstrated. Measurements of
both electroosmotic mobility and selectivity showed that a 2-bilayer and a 20-bilayer PEM
coating could be completely removed from the capillary surface after approximately 3.5
and 9.5 hours, respectively, of continuous exposure to 1.0 M NaOH.
146
The studies discussed earlier verify that the PEM coating has a significant effect on
separations. The next goal is to characterize the coating using various characterization
techniques, such as nuclear magnetic resonance (NMR) spectroscopy, ellipsometry, X-ray
diffraction, Fourier transform infrared (FTIR) spectroscopy, neutron reflectometry,
fluorescence resonance energy transfer (FRET), and near-infrared (NIR) multispectral
imaging spectrometry. Different kinds of microscopic techniques, such as atomic force
microscopy (AFM), scanning electron microscopy (SEM), transmission electron
microscopy (TEM), scanning tunneling microscopy (STM), and near-field scanning optical
microscopy (NSOM) can also be used for the characterization of the coating. In particular,
the NIR multispectral imaging technique, which can simultaneously record the spectral and
spatial information of a sample [1, 2], will be used to study the uniformity and
discontinuity of the coating, and to find the optimized flush times of both PDADMAC and
polymeric surfactant. Then, FRET and some microscopic techniques will be used to
measure the thickness of the PEM coating. FRET is a distance-dependent interaction
between the electronic excited states of two fluorophores. Excitation is transferred from a
donor fluorophore (i.e., fluorescein) to an acceptor fluorophore (i.e.,
tetramethylrhodamine) without emission of a photon. The efficiency of FRET is inversely
proportional to the sixth power of the intermolecular separation.
Although LSCM was able to demonstrate the existence of the coating and the
change in structure after rinsing the capillary with NaOH, it did not provide enough details
regarding the structure of the coating. Therefore, the techniques mentioned above can be
used to provide information about film thickness, interlayer spacings, structure, elemental
content etc [3-14].
147
6.1 References
1. Fischer, M.; Tran, C. D. Anal. Chem. 1999, 71, 2255.
2. Khait, O.; Smirnov, S.; Tran, C. D. Anal. Chem. 2001, 73, 732.
3. Kerimo, J.; Adams, D. M.; Brbara, P. F.; Kaschak, D. M.; Mallouk, T. E. J. Phys. Chem. 1998, 102, 9451.
4. Barberi, R.; Bonvent J. J.; Bartolino, R.; Roeraade, J.; Capelli, L.; Righetti, P. G. J.
Chromatogr. B 1996, 683, 3.
5. Pullen, P. E.; Pesek, J. J.; Matyska, M. T.; Frommer, J. Anal. Chem. 2000, 72, 2751.
6. Mahltig, B.; Müller-Buschbaum, P.; Wolkenhauer, M.; Wunnicke, O.; Wiegand, S.;
Gohy, J.-F.; Jerome, R.; Stamm, M. J. Colloid and Interface Science 2001, 242, 36.
7. Perez-Salas, U. A.; Faucher, K. M.; Majkrzak, C. F.; Berk, N. F.; Krueger, S.; Chakof, E. L. 2003, 19, 7688.
8. Styrkas, D. A.; Bütün, V.; Lu, J. R.; Keddie, J. L.; Armes, S. P. Langmuir 2000, 16,
5980.
9. Oehlke, J.; Birth, P.; Klauschenz, E.; Wiesner, B.; Beyermann, M.; Oksche, A.; Biener, M. Eur. J. Biochem. 2002, 269, 4025.
10. Kharlampieva, E.; Sukhishvili, S. A. Langmuir 2003, 19, 1235.
11. Preisler, J.; Yeung, E. S. Anal. Chem. 1996, 68, 2885.
12. Kaupp, S.; Wätzig, H. J. Chromatogr. A 1997, 781, 55.
13. Kaupp, S.; Steffen, R.; Wätzig, H. J. Chromatogr. A 1996, 744, 93.
14. Advincula, R.; Wang, Y.; Park, M.; Youk, J. H.; Mays, J.; Yang, J.; Kaneko, F.;
Baba, A.; Knoll, W. Polymer Reprints 2001, 42, 281.
148
VITA
Constantina P. Kapnissi was born in Nicosia, Cyprus, on March 22, 1977. She is
the first-born child in the family, and she has three sisters. The fourth one is ten years old.
In June of 1995, she graduated from a public high school. Two months after that she
attended the University of Cyprus. She majored in general chemistry, and she obtained a
Bachelor of Science degree in May, 1999. However, before graduation, in the summer of
1998, she started her undergraduate thesis work at ETH University in Zurich, Switzerland.
After spending a month there, she moved back to Cyprus to continue her work with the
help and direction of her advisor Dr. Epameinondas Leontidis. Her undergraduate thesis
title was “Morphology of Gold Colloids from Cationic Surfactant Solutions.” On July 11,
1999, she got engaged to Andreas Christodoulou, who had already been a student at
Louisiana State University in Baton Rouge, Louisiana. After the engagement, she moved
to United States with him to pursue a degree of Doctor of Philosophy in chemistry at the
same university, under the direction of Dr. Isiah M. Warner. Her dissertation focuses on
the development of packed and open-tubular capillary electrochromatographic methods for
improved achiral and chiral separations of various classes of analytes.
The articles she published, and the patent she submitted from her research work are
listed below:
• Warner, I. M., Kapnissi, C. P., Kamande, M., Valle, B. C., Schlenoff, J. B. Analytical Separations With Polyelectrolyte Layers, Molecular Micelles, or Zwitterionic Polymers, U.S. Patent, Submitted.
• Kapnissi, C. P., Agbaria, R. A., Lowry, M., Geng, L., Warner, I. M. Investigation
of the Stability of Polyelectrolyte Multilayer Coatings Using Laser Scanning Confocal Microscopy and Open-Tubular Capillary Electrochromatography, Submitted for publication.
149
• Kamande, M. W., Zhu, X., Kapnissi-Christodoulou, C. P., Warner, I. M. Chiral Separations Using a Polypeptide, a Polymeric Dipeptide Surfactant, and a Polyelectrolyte Multilayer Coating in Open-Tubular Capillary Electrochromatography, Submitted for publication.
• Kapnissi, C. P., Warner, I. M. Separation of Benzodiazepines by Use of Capillary
Electrochromatography, Journal of Chromatographic Science, 2004, 42, 238-244.
• Zhu, X., Kamande, M. W., Thiam, S., Kapnissi, C. P., Mwongela, S. M., Warner, I. M. Open-Tubular Capillary Electrochromatography/Electrospray Ionization-Mass Spectrometry Using Polymeric Surfactant as a Stationary Phase Coating, Electrophoresis, 2004, 25, 562-568.
• Kapnissi, C. P., Valle, B. C., Warner, I. M. Chiral Separations Using Polymeric
Surfactants and Polyelectrolyte Multilayers in Open-Tubular Capillary Electrochromatography, Analytical Chemistry, 2003, 75, 6097-6104.
• Kapnissi-Christodoulou, C. P., Zhu, X., Warner, I. M. Analytical Separations in
Open-Tubular Capillary Electrochromatography, Electrophoresis, 2003, 24, 3917-3934.
• Kamande, M. W., Kapnissi, C. P., Akbay, C., Zhu, X., Agbaria, R. A., Warner, I.
M. Open-Tubular Capillary Electrochromatography Using a Polymeric Surfactant Coating, Electrophoresis, 2003, 24, 945-951.
• Kapnissi, C. P., Akbay, C., Schlenoff, J. B., Warner, I. M. Analytical Separations
Using Molecular Micelles in Open-Tubular Capillary Electrochromatography, Analytical Chemistry, 2002, 74, 2328-2335.
During her years in graduate school, she presented her work at a number of scientific
conferences including:
• “Chiral Separations Using Polymeric Surfactants and Polyelectrolyte Multilayers in Open-Tubular Capillary Electrochromatography - Morphologic Elucidation of Capillaries Using Laser Scanning Confocal Microscopy” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, Chicago, IL, March 2004. Kapnissi-Christodoulou, C. P., Agbaria, R. A., Lowry, M., Valle, B. C., Geng, L., Warner, I. M.
• “Chiral Separations Using a Polymeric Dipeptide Surfactant in Open-Tubular
Capillary Electrochromatography” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, Chicago, IL, March 2004. Kamande, M. W., Kapnissi-Christodoulou, C. P., Zhu, X., Warner, I. M.
150
• “Chiral Separations Using Polymeric Surfactants and Polyelectrolyte Multilayers in Open-Tubular Capillary Electrochromatography” American Chemical Society Meeting, New York, NY, September 2003. Kapnissi, C. P., Agbaria, R. A., Lowry, M., Valle, B. C., Geng, L., Warner, I. M.
• “Analytical Separations Using Molecular Micelles in Open-Tubular Capillary
Electrochromatography: Morphologic Elucidation of Capillaries Modified by Polyelectrolyte Multilayers” American Chemical Society Meeting, New Orleans, LA, March 2003. Kapnissi, C. P., Agbaria, R. A., Lowry, M., Valle, B. C., Geng, L., Warner, I. M.
• “Separation of Phenols and Benzodiazepines Using Poly (Sodium Undecylenic
Sulfate) and Open-Tubular Capillary Electrochromatography” American Chemical Society Meeting, New Orleans, LA, March 2003. Kamande, M. W., Kapnissi, C. P., Zhu, X., Akbay, C., Agbaria, R. A., Warner, I. M.
• “Morphologic Elucidation of Capillaries Modified by Polyelectrolyte Multilayers
for Analytical Separations” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, Orlando, FL, March 2003. Kapnissi, C. P., Agbaria, R. A., Lowry, M., Valle, B. C., Geng, L., Warner, I. M.
• “Elution Behavior of Hydrophobic Analytes by Use of Various Stationary Phases
in Capillary Electrochromatography” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, Orlando, FL, March 2003. Igbinosun, O. J., Kapnissi, C. P., Kimberly Y. H., Warner, I. M.
• “Open-tubular Capillary Electrochromatography Using a Polymeric Surfactant
Coating” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, Orlando, FL, March 2003. Kamande, M. W., Kapnissi, C. P., Zhu, X., Akbay, C., Agbaria, R. A., Warner, I. M.
• “Open-Tubular Capillary Electrochromatography Using a Polymeric Surfactant
Coating” 16th International Symposium on Microscale Separations and Analysis, San Diego, CA, January 2003. Kamande, M. W., Kapnissi, C. P., Zhu, X., Akbay, C., Warner, I. M.
• “Novel Separations Using Polymeric Surfactants in Open-Tubular Capillary
Electrochromatography” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, New Orleans, LA, March 2002. Kapnissi, C. P., Schlenoff, J. B., Warner, I. M.
• “Separation of Beta-Blockers Using Capillary Electrochromatography” Pittsburgh
Conference of Analytical Chemistry and Applied Spectroscopy, New Orleans, LA, March 2002. Kapnissi, C. P., Igbinosun, O. J., Agbaria, R. A., Warner, I. M.
151
• “Separation of Benzodiazepines by Capillary Electrochromatography” Pittsburgh Conference of Analytical Chemistry and Applied Spectroscopy, New Orleans, LA, March 2001. Kapnissi, C. P., Thiam, S., Warner, I. M.
• “Separation of Benzodiazepines by Capillary Electrochromatography” American
Chemical Society Meeting, New Orleans, LA, December 2000. Kapnissi, C. P., Thiam, S., Warner, I. M.